The project was supported in part by National Institutes of Health Grants RO1 NS099409, NS075534, and CONACYT 2012-037(S) to VP.

1. Preparation
<ol>
	<li>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).</li>
	<li>Prepare 60 mm x 15 mm Petri dishes with apple juice-agar (3% agar and 30% apple juice).</li>
	<li>Prepare fresh yeast paste by mixing dry yeast granules and water. Spread the yeast paste onto the apple agar plates to increase egg laying.</li>
	<li>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.</li>
	<li>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.</li>
	<li>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 &#176;C) and humidity (60%).</li>
	<li>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.</li>
</ol>
2. Collection of Embryos
<ol>
	<li>Carefully pick embryos with a wet paintbrush and place them in a collecting glass dish filled with 1x PBS.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Make a rectangular wax boundary on another glass slide using a wax pen (see Table of Materials).</li>
	<li>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.</li>
	<li>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.</li>
</ol>
3. Recording of Embryos
<ol>
	<li>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).</li>
	<li>Perform live video recording of embryos using suitable software (see Table of Materials) for about 1-2 h&#160;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).</li>
</ol>
4. Analysis of the Recordings
<ol>
	<li>Export the recorded video directly into Image J for further analyses (e.g., as AVI files).</li>
	<li>In ImageJ, crop the video recordings to the size of individual embryos by drawing a box around each embryo and then clicking on Image &#62; Crop. This greatly reduces the size of video files without affecting its resolution and facilitates their analysis.</li>
	<li>Rotate cropped images to achieve vertical position of the embryo midline relative to the screen, by clicking on Image &#62; Transform &#62; Rotate. 
	NOTE: Selecting Preview during this process will provide guidance for rotation, showing gridlines to ensure vertical position of the midline.</li>
	<li>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&#160;&#62;&#160;Set Scale, and then entering a conversion of pixels to distances, e.g., micrometers
	<ol>
		<li>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 &#62; Tools &#62; ROI manager and record this position as slice number-y coordinate-x coordinate by drawing a box of approximately 7 &#181;m by&#160;7 &#181;m 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.</li>
		<li>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 &#181;m using a known scale of images. Alternatively, measure the distance in &#181;m in a single step by clicking on Analyze &#62; 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 &#181;m.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Quantitative analysis of embryonic muscle contractions:
	<ol>
		<li>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.</li>
		<li>Use the recording of fluorescent readout and draw a region of interest (e.g., a box of 15 &#181;m to 45 &#181;m [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 &#62; Measure to record the average fluorescent intensity of each region of interest selected for each frame of the video.</li>
		<li>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.</li>
		<li>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 &#62; 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.</li>
		<li>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).</li>
		<li>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:&#160;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.</li>
		<li>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.</li>
		<li>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).</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

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...

Peristaltic muscle contraction is a rhythmic motor behavior similar to walking and swimming in humans1,2,3. 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 direction1,2. 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 muscles4,5. 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 proteins8,9. Genetic defects in POM cause congenital muscular dystrophies (CMD) in humans10,11,12. 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 (&#34;rotation&#34;) 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.

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

live imaging and analysis of muscle contractions in drosophila embryo

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.

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...

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Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

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

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Coordinated muscle contractions are a form of rhythmic behavior seen early during development in Drosophila embryos. Neuronal sensory feedback circuits ...

live-imaging-and-analysis-of-muscle-contractions-in-drosophila-embryo

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6.0K Views.  Texas A&M University. Here, we present a method to record embryonic muscle contractions in Drosophila embryos in a non-invasive and detail-oriented manner.

Video: Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

Live-imaging of the Drosophila Pupal Eye

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track during live cell imaging. These processes include: retinal rotation, cell growth and organismal movement. Additionally, irregularities in the topology of the epithelium, including subtle bumps and folds often introduced as the pupa is prepared for imaging, make it challenging to acquire in-focus images of more than a few ommatidia in a single focal plane. The workflow outlined here remedies these issues, allowing easy analysis of cellular processes during Drosophila pupal eye development. Appropriately-staged pupae are arranged in an imaging rig that can be easily assembled in most laboratories. Ubiquitin-DE-Cadherin:GFP and GMR-GAL4&#8211;driven UAS-&#945;-catenin:GFP are used to visualize cell boundaries in the eye epithelium 1-3. After deconvolution is applied to fluorescent images captured at multiple focal planes, maximum projection images are generated for each time point and enhanced using image editing software. Alignment algorithms are used to quickly stabilize superfluous motion, making individual cell behavior easier to track.

Live-imaging of the Drosophila Pupal Eye

This protocol presents an efficient method for imaging the live Drosophila pupal eye neuroepithelium. This method compensates for tissue movement and uneven topology, enhances visualization of cell boundaries through the use of multiple GFP-tagged junction proteins, and uses an easily-assembled imaging rig.

Inherent processes of Drosophila pupal development can shift and distort the eye epithelium in ways that make individual cell behavior difficult to track ...

Long-term Live Imaging of Drosophila Eye Disc

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have been used to study many biological processes, such as cell proliferation, differentiation, growth, migration, apoptosis, competition, cell-cell signaling, and compartmental boundary formation. However, methods for the long-term ex vivo culture and live imaging of the imaginal discs have not been satisfactory, despite many efforts. Recently, we developed a method for the long-term ex vivo culture and live imaging of imaginal discs for up to 18 h. In addition to using a high insulin concentration in the culture medium, a low-melting agarose was also used to embed the disc to prevent it from drifting during the imaging period. This report uses the eye-antennal discs as an example. Photoreceptor R3/4-specific m&#948;0.5-Ga4 expression was followed to demonstrate that photoreceptor differentiation and ommatidial rotation can be observed during a 10 h live imaging period. This is a detailed protocol describing this simple method.

Long-term Live Imaging of Drosophila Eye Disc

This protocol describes a detailed method for the long-term ex vivo culture and live imaging of a Drosophila imaginal disc. It demonstrates photoreceptor differentiation and ommatidial rotation within the 10 h period of live imaging of the eye disc. The protocol is simple and does not require expensive setup.

Live imaging provides the ability to continuously track dynamic cellular and developmental processes in real time. Drosophila larval imaginal discs have ...

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from animals to humans. Therefore, the powerful Drosophila model system can be used to study concepts of muscle-tendon development that can also be applied to human muscle biology. Here, we describe in detail how morphogenesis of the adult muscle-tendon system can be easily imaged in living, developing Drosophila pupae. Hence, the method allows investigating proteins, cells and tissues in their physiological environment. In addition to a step-by-step protocol with helpful tips, we provide a comprehensive overview of fluorescently tagged marker proteins that are suitable for studying the muscle-tendon system. To highlight the versatile applications of the protocol, we show example movies ranging from visualization of long-term morphogenetic events &#8211; occurring on the time scale of hours and days &#8211; to visualization of short-term dynamic processes like muscle twitching occurring on time scale of seconds. Taken together, this protocol should enable the reader to design and perform live-imaging experiments for investigating muscle-tendon morphogenesis in the intact organism.

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Here, we present an easy-to-use and versatile method to perform live imaging of developmental processes in general and muscle-tendon morphogenesis in particular in living Drosophila pupae.

Muscles together with tendons and the skeleton enable animals including humans to move their body parts. Muscle morphogenesis is highly conserved from ...

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of the Drosophila melanogaster syncytial embryo for studying mitosis1. Drosophila has useful genetics with a sequenced genome, and it can be easily maintained and manipulated. Many mitotic mutants exist, and transgenic flies expressing functional fluorescently (e.g. GFP) - tagged mitotic proteins have been and are being generated. Targeted gene expression is possible using the GAL4/UAS system2. 

The Drosophila early embryo carries out multiple mitoses very rapidly (cell cycle duration, &asymp;10 min). It is well suited for imaging mitosis, because during cycles 10-13, nuclei divide rapidly and synchronously without intervening cytokinesis at the surface of the embryo in a single monolayer just underneath the cortex. These rapidly dividing nuclei probably use the same mitotic machinery as other cells, but they are optimized for speed; the checkpoint is generally believed to not be stringent, allowing the study of mitotic proteins whose absence would cause cell cycle arrest in cells with a strong checkpoint. Embryos expressing GFP labeled proteins or microinjected with fluorescently labeled proteins can be easily imaged to follow live dynamics (Fig. 1). In addition, embryos can be microinjected with function-blocking antibodies or inhibitors of specific proteins to study the effect of the loss or perturbation of their function3. These reagents can diffuse throughout the embryo, reaching many spindles to produce a gradient of concentration of inhibitor, which in turn results in a gradient of defects comparable to an allelic series of mutants. Ideally, if the target protein is fluorescently labeled, the gradient of inhibition can be directly visualized4. It is assumed that the strongest phenotype is comparable to the null phenotype, although it is hard to formally exclude the possibility that the antibodies may have dominant effects in rare instances, so rigorous controls and cautious interpretation must be applied. Further away from the injection site, protein function is only partially lost allowing other functions of the target protein to become evident.

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

This protocol describes the use of microinjection and high resolution imaging in the Drosophila melanogaster syncytial embryo to study mitosis.

This protocol describes the use of the Drosophila melanogaster syncytial embryo  for studying mitosis1.  Drosophila has useful genetics with a sequenced ...

Biology

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ellipsoid shape during development. This developmental process is called oogenesis and is crucial to generating functional mature eggs to secure the next fly generation. For these reasons, egg chambers have served as an ideal and relevant model to understand animal organ development.
Several in vitro culturing protocols have been developed, but there are several disadvantages to these protocols. One involves the application of various covers that exert an artificial pressure on the imaged egg chambers in order to immobilize them and to increase the imaged acquisition plane of the circumferential surface of the analyzed egg chambers. Such an approach may negatively influence the behavior of the thin actomyosin machinery that generates the power to rotate egg chambers around their longer axis.
Thus, to overcome this limitation, we culture Drosophila egg chambers freely in the media in order to reliably analyze actomyosin machinery along the circumference of egg chambers. In the first part of the protocol, we provide a manual detailing how to analyze the actomyosin machinery in a limited acquisition plane at the local cellular scale (up to 15 cells). In the second part of the protocol, we provide users with a new Fiji-based plugin that allows the simple extraction of a defined thin layer of the egg chambers&#8217; circumferential surface. The following protocol then describes how to analyze actomyosin signals at the tissue scale (&#62;50 cells). Finally, we pinpoint the limitations of these approaches at both the local cellular and tissue scales and discuss its potential future development and possible applications.

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

This protocol provides a Fiji-based, user-friendly methodology along with straightforward instructions explaining how to reliably analyze actomyosin behavior in individual cells and curved epithelial tissues. No programming skills are required to follow the tutorial; all steps are performed in a semi-interactive manner using the graphical user interface of Fiji and associated plugins.

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ...

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we present a protocol for the mounting of Drosophila melanogaster embryos and subsequent live imaging of fluorescently labeled hemocytes, the embryonic macrophages of this organism. Using the Gal4-uas system1 we drive the expression of a variety of genetically encoded, fluorescently tagged markers in hemocytes to follow their developmental dispersal throughout the embryo. Following collection of embryos at the desired stage of development, the outer chorion is removed and the embryos are then mounted in halocarbon oil between a hydrophobic, gas-permeable membrane and a glass coverslip for live imaging. In addition to gross migratory parameters such as speed and directionality, higher resolution imaging coupled with the use of fluorescent reporters of F-actin and microtubules can provide more detailed information concerning the dynamics of these cytoskeletal components.

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Drosophila hemocytes disperse over the entirety of the developing embryo. This protocol demonstrates how to mount and image these migrations using embryos with fluorescently labelled hemocytes.

Many studies address cell migration using in vitro methods, whereas the physiologically relevant environment is that of the organism itself. Here we ...

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A remarkable example is given by sensory epithelia, where cells of the same or distinct identities are brought together via cell-cell adhesion showing highly organized planar patterns. Cells align to one another in the same direction and display equivalent polarity over large distances. This organization of the mature epithelia is established over the course of morphogenesis. To understand how the planar arrangement of the mature epithelia is achieved, it is crucial to track cell orientation and growth dynamics with high spatiotemporal fidelity during development in vivo. Robust analytical tools are also essential to identify and characterize local-to-global transitions. The Drosophila pupa is an ideal system to evaluate oriented cell shape changes underlying epithelial morphogenesis. The pupal developing epithelium constitutes the external surface of the immobile body, allowing long-term imaging of intact animals. The protocol described here is designed to image and analyze cell behaviors at both global and local levels in the pupal abdominal epidermis as it grows. The methodology described can be easily adapted to the imaging of cell behaviors at other developmental stages, tissues, subcellular structures, or model organisms.

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

This protocol is designed for the imaging and analysis of the dynamics of cell orientation and tissue growth in the Drosophila abdominal epithelia as the fruit fly undergoes metamorphosis. The methodology described here can be applied to the study of different developmental stages, tissues, and subcellular structures in Drosophila or other model organisms.

Within multicellular organisms, mature tissues and organs display high degrees of order in the spatial arrangements of their constituent cells. A ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Neuroscience

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and computational modeling to classify and compare biological processes. Although cell culture model systems are great for high content imaging, high throughput studies of cell morphology suggest that ex vivo cultures are limited in recapitulating the morphological complexity found in cells within living organisms. As such, there is a need for a scalable high throughput model system to image living cells within an intact organism. Described here is a protocol for using a high content image analyzer to simultaneously acquire multiple time-lapse videos of embryonic Drosophila melanogaster development during the syncytial blastoderm stage. The syncytial blastoderm has traditionally served as a great in vivo model for imaging biological events; however, obtaining a significant number of experimental replicates for quantitative and high-throughput approaches has been labor intensive and limited by the imaging of a single embryo per experimental repeat. Presented here is a method to adapt imaging and microinjection approaches to suit a high content imaging system, or any inverted microscope capable of automated multipoint acquisition. This approach enables the simultaneous acquisition of 6-12 embryos, depending on desired acquisition factors, within a single imaging session.

Presented here is a protocol to microinject and simultaneously image multiple Drosophila embryos during embryonic development using a plate-based, high content imager.

Modern approaches in quantitative live cell imaging have become an essential tool for exploring cell biology, by enabling the use of statistics and ...

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal lines have enabled biological processes to be studied in the context of a living, and in some instances even behaving, organism. In this protocol we will describe how to anesthetize intact Drosophila larvae, using the volatile anesthetic desflurane, to follow the development and plasticity of synaptic populations at sub-cellular resolution1-3. While other useful methods to anesthetize Drosophila melanogaster larvae have been previously described4,5,6,7,8, the protocol presented herein demonstrates significant improvements due to the following combined key features: (1) A very high degree of anesthetization; even the heart beat is arrested allowing for lateral resolution of up to 150 nm1, (2) a high survival rate of &gt; 90% per anesthetization cycle, permitting the recording of more than five time-points over a period of hours to days2 and (3) a high sensitivity enabling us in 2 instances to study the dynamics of proteins expressed at physiological levels. In detail, we were able to visualize the postsynaptic glutamate receptor subunit GluR-IIA expressed via the endogenous promoter1 in stable transgenic lines and the exon trap line FasII-GFP1. (4) In contrast to other methods4,7 the larvae can be imaged not only alive, but also intact (i.e. non-dissected) allowing observation to occur over a number of days1. The accompanying video details the function of individual parts of the in vivo imaging chamber2,3, the correct mounting of the larvae, the anesthetization procedure, how to re-identify specific positions within a larva and the safe removal of the larvae from the imaging chamber.

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution

This protocol describes a reliable method for anesthetization and imaging of intact Drosophila melanogaster larvae. We have utilized the volatile anesthetic desflurane to allow for repetitive imaging at sub-cellular resolution and re-identification of structures for up to a few days1.

Recent improvements in optical imaging, genetically encoded fluorophores and genetic tools allowing efficient establishment of desired transgenic animal ...

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

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Live-imaging of the Drosophila Pupal Eye

Long-term Live Imaging of Drosophila Eye Disc

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Microinjection Techniques for Studying Mitosis in the Drosophila melanogaster Syncytial Embryo

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Imaging and Analysis of Tissue Orientation and Growth Dynamics in the Developing Drosophila Epithelia During Pupal Stages

Live Imaging of Drosophila Larval Neuroblasts

Drosophila Embryo Preparation and Microinjection for Live Cell Microscopy Performed using an Automated High Content Analyzer

In vivo Imaging of Intact Drosophila Larvae at Sub-cellular Resolution