Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

1. Preparation

1. Prepare a fly cage by making approximately 50 holes in a 100 mL capacity tri-corner plastic beaker using a hot 25 G needle (see Table of Materials).
2. Prepare 60 mm x 15 mm Petri dishes with apple juice-agar (3% agar and 30% apple juice).
3. Prepare fresh yeast paste by mixing dry yeast granules and water. Spread the yeast paste onto the apple agar plates to increase egg laying.
4. Anaesthetize about 50-60 flies (use approximately equal numbers of males and females) on CO₂ and put them in the fly cage.
   NOTE: Using an increased proportion of females (up to ~2:1 ratio of females:males) may help increase the amount of laid eggs for some genotypes.
5. Attach an apple juice-agar Petri dish with yeast paste to the fly cage tightly and seal it with modeling clay. Make sure it is sealed at all corners.
6. Wait until flies wake up from anesthesia and then invert the cage such that the Petri dish is now at the bottom. Put the cage into an incubator with controlled temperature (25 °C) and humidity (60%).
7. Allow flies to lay eggs for 2-3 h, replace the apple plate with a fresh one, and let the plate with eggs age for 19-20 h in an incubator.
   NOTE: Prior to the above step, flies must be synchronized to facilitate collection of stage 17e-f (19-21 h AEL) embryos. This can be achieved by transferring flies to a cage with a fresh yeast-apple juice-agar plate 3-4 times for 12 h (once every 3-4 h). Keeping flies at controlled circadian light environment (LD cycle) can also help with collecting a synchronized population of embryos, but this was not essential in our experiments.

2. Collection of Embryos

1. Carefully pick embryos with a wet paintbrush and place them in a collecting glass dish filled with 1x PBS.
2. Select the embryos that have their tracheae filled with air. Air-filled tracheae indicate that embryos have reached Stage 17, and their peristaltic muscle contractions should have begun. Tracheae become clearly visible when they are filled with air, which can serve as a marker for Stage 17.
3. Place an apple juice agar slab on a glass slide and carefully transfer embryos from PBS to the slab. Line up the embryos with their ventral side up.
   NOTE: Dorsal and ventral sides of embryos can be distinguished by the position of dorsal appendages on the eggshell.
4. Make a rectangular wax boundary on another glass slide using a wax pen (see Table of Materials).
5. Place a double-sided sticky tape within that boundary and gently pick up the embryos by lowering this slide onto the agar slab. Apply gentle pressure to ensure that embryos stick to the tape well, with their dorsal side up. If necessary, embryos can be rolled on the tape to correct their orientation. Do all manipulations while monitoring embryos under a dissection microscope.
6. Cover embryos with 1x PBS for live imaging of muscle contractions.
   NOTE: Some procedures described above are related to basic Drosophila techniques used in many studies. More detailed descriptions of common Drosophila techniques can be found elsewhere¹⁴.

3. Recording of Embryos

1. Perform live imaging of mounted embryos on an epiflourescence microscope with a time-lapse function and a digital camera with suitable emission filters (see Table of Materials) using a 10x water immersion objective lens.
   NOTE: Here we used embryos expressing GFP in muscles. Other fluorescent markers with suitable excitation light and emission filter sets can also be used (e.g., for tdTomato detection, one can use a Chroma ET-561 filter set for excitation and emission around optimal 554 nm and 581 nm, respectively).
2. Perform live video recording of embryos using suitable software (see Table of Materials) for about 1-2 h with an acquisition rate of 4 frames/s.
   NOTE: To analyze rolling of the developing embryo within its shell, embryos without expression of fluorescent markers can be used. To this end, regular transmitted light illumination without spectral filters is applied to visualize embryo motion within the shell (See Movie 2).

4. Analysis of the Recordings

1. Export the recorded video directly into Image J for further analyses (e.g., as AVI files).
2. In ImageJ, crop the video recordings to the size of individual embryos by drawing a box around each embryo and then clicking on Image > Crop. This greatly reduces the size of video files without affecting its resolution and facilitates their analysis.
3. Rotate cropped images to achieve vertical position of the embryo midline relative to the screen, by clicking on Image > Transform > Rotate.
   NOTE: Selecting プレビュー during this process will provide guidance for rotation, showing gridlines to ensure vertical position of the midline.
4. Quantitative analysis of embryo rolling:
   NOTE: For distance analyses, first confirm that your images include scale information. Image scale can be added by selecting Analyze > Set Scale, and then entering a conversion of pixels to distances, e.g., micrometers
   1. Mark the position of one or both tracheae in the first frame of the video at a point midway between posterior and anterior ends. Click on Analyze > Tools > ROI manager and record this position as slice number-y coordinate-x coordinate by drawing a box of approximately 7 µm by 7 µm around it and typing t on the keyboard. Ensure that when typing t, a region of interest is selected on the video. Alternatively, select the Add (t) tab on the ROI manager to record the position of trachea instead of typing the command.
      NOTE: The region of interest can vary in shape or size depending on the embryonic region or developmental event being studied.
   2. Mark the position of the same area of the trachea after each peristaltic contraction. Measure the distance from the pre-contraction position to the post-contraction position by drawing a line connecting the centers of each box and typing m on the keyboard. Convert the distance to µm using a known scale of images. Alternatively, measure the distance in µm in a single step by clicking on Analyze > Set Scale and enter the known pixel-to-micron conversion factor to yield a report in microns.
      NOTE: A distance in pixels can be entered together with its corresponding distance in µm.
   3. Correlate the distance and direction of each rolling event with the direction of muscle contraction propagation in at least 8 embryos for statistically significant differences.
5. Quantitative analysis of embryonic muscle contractions:
   1. Use embryos expressing fluorescent markers in muscles (e.g., we used transgenic flies expressing a fusion construct of Myosin Heavy Chain promoter and GFP called MHC-GFP⁵) to analyze muscle contraction parameters such as contraction amplitude.
   2. Use the recording of fluorescent readout and draw a region of interest (e.g., a box of 15 µm to 45 µm [HXW]) centered on the muscles (which are clearly visible due to the presence of fluorescent marker) of a particular body segment, and select the Add (t) tab on the ROI manager to record the position of the ROI. Click on ROI manager > Measure to record the average fluorescent intensity of each region of interest selected for each frame of the video.
   3. Move the box to the centers of other body segments of interest and click on Add (t) in the ROI manager to record their positions. This will give regions of interest of identical size in all body segments to be analyzed. Select at least one posterior, one medial, and one anterior segment, e.g., A7, A4, and T2, respectively.
   4. In the ROI manager, select all regions of interest recorded as slice number-y coordinate-x coordinate (e.g., by selecting while holding Ctrl) and click on 詳細 > Multi measure to measure the mean fluorescent intensity of each region of interest for all frames of the video, and report each measurement in a table. Each region of interest is a column of the table, and each frame is a row. Transfer the table to a spreadsheet program for further analyses.
   5. Plot a graph with frame number on the x-axis and mean fluorescent intensity on the y-axis. Frame number can be converted to time using the frame rate (4 frames/s) of the video (Figure 1A).
   6. Determine muscle contraction amplitude by estimating the increase in GFP fluorescence intensity relative to the baseline. Muscle contractions increase the GFP intensity as they bring more GFP into the vicinity of the focal area as more muscles get pulled in during these contractions (Movie 1)⁷. Establish a baseline fluorescence as the average intensity between contraction waves. Normalize GFP intensity to the baseline by dividing every ROI intensity value by the baseline intensity.
      NOTE: Each profile has a different baseline fluorescence, as there may be different expression levels in different muscle segments.
      NOTE: One potential complication is that the GFP fluorescence may change over time due to photo bleaching. This can be resolved by monitoring changes in fluorescence baseline and using a sufficient sample size for wave analyses (we normally use sets of 10 fluorescent waves and confirm that the baseline is approximately constant by taking an average of only those peak minima as baseline that have decreased in fluorescence by 10% or less relative to the initial minima peak). A pulse-LED illumination may be also applied to mitigate that problem¹⁶.
   7. Compare muscle contractions on left and right sides of the embryo by analyzing peak intensities on both sides of the embryo for same segments. Use contraction amplitude and time of peak intensities to examine differences in extent and timing of peristaltic muscle contraction waves propagating along both sides of the embryo.
   8. Compare normalized intensity of GFP at different segments (e.g., at anterior, medial and posterior regions) during muscle contraction wave propagation to examine changes in the contraction as the wave propagates. This analysis also determines the direction of the wave (i.e., whether it propagates toward anterior or posterior regions of the embryo).