Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Podsumowanie

Here, we present a method to record embryonic muscle contractions in Drosophila embryos in a non-invasive and detail-oriented manner.

Streszczenie

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits are required to control this behavior. Failure to produce the rhythmic pattern of contractions can be indicative of neurological abnormalities. We previously found that defects in protein O-mannosylation, a posttranslational protein modification, affect the axon morphology of sensory neurons and result in abnormal coordinated muscle contractions in embryos. Here, we present a relatively simple method for recording and analyzing the pattern of peristaltic muscle contractions by live imaging of late stage embryos up to the point of hatching, which we used to characterize the muscle contraction phenotype of protein O-mannosyltransferase mutants. Data obtained from these recordings can be used to analyze muscle contraction waves, including frequency, direction of propagation and relative amplitude of muscle contractions at different body segments. We have also examined body posture and taken advantage of a fluorescent marker expressed specifically in muscles to accurately determine the position of the embryo midline. A similar approach can also be utilized to study various other behaviors during development, such as embryo rolling and hatching.

Wprowadzenie

Peristaltic muscle contraction is a rhythmic motor behavior similar to walking and swimming in humans^1,2,3. Embryonic muscle contractions seen in Drosophila late stage embryos represent an example of such a behavior. Drosophila is an excellent model organism to study various developmental processes because embryonic development in Drosophila is well characterized, relatively short, and easy to monitor. The overall goal of our method is to carefully record and analyze the wavelike pattern of contraction and relaxation of embryonic muscles. We used a simple, non-invasive approach that offers a detailed visualization, recording and analysis of muscle contractions. This method can also be potentially used to study other in vivo processes, such as embryonic rolling seen in late stage embryos just prior to hatching. In previous studies, embryonic muscle contractions were mostly analyzed in terms of frequency and direction^1,2. In order to estimate the relative extent of contractions as they progress along the body axis in the anterior or posterior direction, we have used embryos expressing GFP specifically in muscles. This analysis provides a more quantitative way to analyze muscle contractions and to reveal how body posture in embryos is maintained during series of peristaltic waves of muscle contractions.

Peristaltic muscle contractions are controlled by central pattern generator (CPG) circuits and communications between neurons of the peripheral nervous system (PNS), the central nervous system (CNS), and muscles^4,5. Failure to produce normal peristaltic muscle contractions can lead to defects such as failure to hatch² and abnormal larval locomotion⁶ and can be indicative of neurological abnormalities. Live imaging of peristaltic waves of muscle contraction and detailed analysis of contraction phenotypes can help uncover pathogenic mechanisms associated with genetic defects affecting muscles and neural circuits involved in locomotion. We recently used that approach to investigate mechanisms that result in a body posture torsion phenotype of protein O-mannosyltransferase (POMT) mutants⁷.

Protein O-mannosylation (POM) is a special type of posttranslational modification, where a mannose sugar is added to serine or threonine residues of secretory pathway proteins^8,9. Genetic defects in POM cause congenital muscular dystrophies (CMD) in humans^10,11,12. We investigated the causative mechanisms of these diseases using Drosophila as a model system. We found that embryos with mutations in Drosophila protein O-mannosyltransferase genes POMT1 and POMT2 (a.k.a. rotated abdomen (rt) and twisted (tw)) show a displacement ("rotation") of body segments, which results in an abnormal body posture⁷. Interestingly, this defect coincided with the developmental stage when peristaltic muscle contractions become prominent⁷.

Since abnormal body posture in POM mutant embryos arises when musculature and epidermis are already formed and peristaltic waves of coordinated muscle contractions have started, we hypothesized that abnormal body posture could be a result of abnormal muscle contractions rather than a defect in muscle or/and epidermis morphology⁷. CMDs can be associated with abnormal muscle contractions and posture defects¹³, and thus the analysis of the posture phenotype in Drosophila POMT mutants may elucidate pathological mechanisms associated with muscular dystrophies. In order to investigate the relationship between the body posture phenotype of Drosophila POMT mutants and possible abnormalities in peristaltic waves of muscle contractions, we decided to analyze muscle contractions in detail using a live imaging approach.

Our analysis of peristaltic contraction waves in Drosophila embryos revealed two distinct contraction modes, designated as type 1 and type 2 waves. Type 1 waves are simple waves propagating from anterior to posterior or vice versa. Type 2 waves are biphasic waves that initiate at the anterior end, propagate halfway in the posterior direction, momentarily halt, forming a temporal static contraction, and then, during the second phase, are swept by a peristaltic contraction that propagates forward from the posterior end. Wild-type embryos normally generate a series of contractions that consists of approximately 75% type 1 and 25% type 2 waves. In contrast, POMT mutant embryos generate type 1 and type 2 waves at approximately equal relative frequencies.

Our approach can provide detailed information for quantitative analysis of muscle contractions and embryo rolling⁷. This approach could also be adapted for analyses of other behaviors involving muscle contractions, such as hatching and crawling.

Protokół

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 Preview 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 More > 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).

Wyniki

Normal peristaltic muscle contractions are shown in a WT (wild-type, Canton-S) embryo in Movie 1. The average frequency of peristaltic waves of muscle contractions in our analysis was 47 contractions per hour and the average amplitude was 60% above baseline for WT embryos. Embryo rolling is shown for a WT embryo in Movie 2, with the white arrow marking the initial position of a trachea and a black arrow depicting the position of a dorsal appendage. The dorsal appendage (exterio...

Dyskusje

Our method provides a quantitative way to analyze important embryo behaviors during development, such as peristaltic muscle contraction waves, including wave periodicity, amplitude and pattern, as well as wave effect on embryo rolling and posture. This can be useful in analyses of different mutants to study the role of specific genes in regulating these and other behaviors during embryonic development. We have used changes in muscle-specific GFP marker intensity to analyze muscle contraction amplitude, frequency and dire...

Materiały

Name	Company	Catalog Number	Comments
Digital camera	Hamamatsu CMOS ORCA-Flash 4.0	C13440-20CU	With different emission filters
Forceps	FST Dumont	11254-20	Tip Dimensions 0.05 mm x 0.01 mm
LED	X-cite BDX (Excelitas)	XLED1
Microscope	Carl Ziess Examiner D1	491405-0005-000	Epiflourescence with time lapse
Needle	BD	305767	25 G x 1-1/2 inch
Paintbrush	Contemporary crafts		Any paintbrush will work
Petri dishes	VWR	25384-164	60 mm x 15 mm
Software	HCImage Live
Thread Zap Wax pen	Thread Zap II (by BeadSmith)(Amazon)	TZ1300	Burner Tool
Tricorner plastic beaker	VWR	25384-152	100 mL