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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F., Fiala, A., Bate, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+development+of+motor+coordination+in+Drosophila+embryos.">The development of motor coordination in Drosophila embryos.</a> Development. 135, 3707-3717 (2008).</li><li>Song, W., Onishi, M., Jan, L. Y., Jan, Y. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Peripheral+multidendritic+sensory+neurons+are+necessary+for+rhythmic+locomotion+behavior+in+Drosophila+larvae.">Peripheral multidendritic sensory neurons are necessary for rhythmic locomotion behavior in Drosophila larvae.</a> Proceedings of National Academy of Science of the United States of America. 104, 5199-5204 (2007).</li><li>Hughes, C. L., Thomas, J. 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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 border="0" cellpadding="0" cellspacing="0" style="width:771px;" width="771">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Digital camera</td>
			<td style="width:155px;">Hamamatsu CMOS ORCA-Flash 4.0</td>
			<td style="width:119px;">C13440-20CU</td>
			<td style="width:283px;">With different emission filters</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Forceps</td>
			<td style="width:155px;">FST Dumont</td>
			<td style="width:119px;">11254-20</td>
			<td>Tip Dimensions 0.05 mm x 0.01 mm</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">LED</td>
			<td style="width:155px;">X-cite BDX (Excelitas)</td>
			<td style="width:119px;">XLED1</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Microscope</td>
			<td style="width:155px;">Carl Ziess Examiner D1</td>
			<td style="width:119px;">491405-0005-000</td>
			<td>Epiflourescence with time lapse</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Needle</td>
			<td style="width:155px;">BD&#160;</td>
			<td align="right" style="width:119px;">305767</td>
			<td>25 G x 1-1/2 in</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Paintbrush</td>
			<td style="width:155px;">Contemporary crafts</td>
			<td style="width:119px;"></td>
			<td>Any paintbrush will work</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Petri dishes</td>
			<td style="width:155px;">VWR</td>
			<td style="width:119px;">25384-164</td>
			<td>60 mm x 15 mm</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:215px;">Software</td>
			<td style="width:155px;">HCImage Live</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:215px;">Thread Zap Wax pen</td>
			<td style="width:155px;">Thread Zap II (by BeadSmith)(Amazon)</td>
			<td style="width:119px;">TZ1300</td>
			<td>Burner Tool</td>
		</tr>
		<tr height="23">
			<td height="23" style="height:23px;width:215px;">Tricorner plastic beaker</td>
			<td style="width:155px;">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...

Watch this Scientific Journal Video about Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo at JoVE.com

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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Developmental Biology

5.9K 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 ...

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

The zebrafish model is an emerging system for the study of neuromuscular disorders. In the study of neuromuscular diseases, the integrity of the muscle membrane is a critical disease determinant. To date, numerous neuromuscular conditions display degenerating muscle fibers with abnormal membrane integrity; this is most commonly observed in muscular dystrophies. Evans Blue Dye (EBD) is a vital, cell permeable dye that is rapidly taken into degenerating, damaged, or apoptotic cells; in contrast, it is not taken up by cells with an intact membrane. EBD injection is commonly employed to ascertain muscle integrity in mouse models of neuromuscular diseases. However, such EBD experiments require muscle dissection and/or sectioning prior to analysis. In contrast, EBD uptake in zebrafish is visualized in live, intact preparations. Here, we demonstrate a simple and straightforward methodology for performing EBD injections and analysis in live zebrafish. In addition, we demonstrate a co-injection strategy to increase efficacy of EBD analysis. Overall, this video article provides an outline to perform EBD injection and characterization in zebrafish models of neuromuscular disease.

In this study, we describe a straightforward method to perform Evans Blue Dye (EBD) analysis on zebrafish larvae. This technique is a powerful tool for the characterization of skeletal muscle integrity and delineation of zebrafish models of muscular dystrophy, and is a valuable method for the development of novel therapeutics.

The zebrafish model is an emerging system for the study of neuromuscular disorders.  In the study of neuromuscular diseases, the integrity of the muscle ...

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin condensation, microtubule-kinetochore attachment, chromosome segregation, and cytokinesis in a small time frame. Errors in the delicate process can result in human disease, including birth defects and cancer. Traditional approaches investigating human mitotic disease states often rely on cell culture systems, which lack the natural physiology and developmental/tissue-specific context advantageous when studying human disease. This protocol overcomes many obstacles by providing a way to visualize, with high resolution, chromosome dynamics in a vertebrate system, the zebrafish. This protocol will detail an approach that can be used to obtain dynamic images of dividing cells, which include: in vitro transcription, zebrafish breeding/collecting, embryo embedding, and time-lapse imaging. Optimization and modifications of this protocol are also explored. Using H2A.F/Z-EGFP (labels chromatin) and mCherry-CAAX (labels cell membrane) mRNA-injected embryos, mitosis in AB wild-type, auroraBhi1045, and esco2hi2865 mutant zebrafish is visualized. High resolution live imaging in zebrafish allows one to observe multiple mitoses to statistically quantify mitotic defects and timing of mitotic progression. In addition, observation of qualitative aspects that define improper mitotic processes (i.e., congression defects, missegregation of chromosomes, etc.) and improper chromosomal outcomes (i.e., aneuploidy, polyploidy, micronuclei, etc.) are observed. This assay can be applied to the observation of tissue differentiation/development and is amenable to the use of mutant zebrafish and pharmacological agents. Visualization of how defects in mitosis lead to cancer and developmental disorders will greatly enhance understanding of the pathogenesis of disease.

Mitosis is critical to every living organism and defects often lead to cancer and developmental disorders. Using this imaging protocol and zebrafish as a model system, researchers can visualize mitosis in a live vertebrate organism and the multitude of defects that arise when mitotic processes are defective.

Mitosis is critical for organismal growth and differentiation. The process is highly dynamic and requires ordered events to accomplish proper chromatin ...

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 Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; for example, how the zygote polarizes to establish the body axis and how the embryo specifies various cell fates during organ formation. This manuscript describes an in vitro ovule culture method to perform live-cell imaging of developing zygotes and embryos of Arabidopsis thaliana. The optimized cultivation medium allows zygotes or early embryos to grow into fertile plants. By combining it with a poly(dimethylsiloxane) (PDMS) micropillar array device, the ovule is held in the liquid medium in the same position. This fixation is crucial to observe the same ovule under a microscope for several days from the zygotic division to the late embryo stage. The resulting live-cell imaging can be used to monitor the real-time dynamics of zygote polarization, such as nuclear migration and cytoskeleton rearrangement, and also the cell division timing and cell fate specification during embryo patterning. Furthermore, this ovule cultivation system can be combined with inhibitor treatments to analyze the effects of various factors on embryo development, and with optical manipulations such as laser disruption to examine the role of cell-cell communication.

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

This manuscript describes an in vitro ovule cultivation method that enables live-cell imaging of Arabidopsis zygotes and embryos. This method is utilized to visualize the intracellular dynamics during zygote polarization and the cell fate specification in developing embryos.

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and differentiation. Typically, pHi dynamics are determined in cultured cells, which are amenable to measuring and experimentally manipulating pHi. However, the recent development of new tools and methodologies has made it possible to study pHi dynamics within intact, live tissue. For Drosophila research, one important development was the generation of a transgenic line carrying a pHi biosensor, mCherry::pHluorin. Here, we describe a protocol that we routinely use for imaging live Drosophila ovarioles to measure pHi in the epithelial follicle stem cell (FSC) lineage in mCherry::pHluorin transgenic wild type lines; however, the methods described here can be easily adapted for other tissues, including the wing discs and eye epithelium. We describe techniques for expressing mCherry::pHluorin in the FSC lineage, maintaining ovarian tissue during live imaging, and acquiring and analyzing images to obtain pHi values.

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

We provide a protocol for imaging intracellular pH of an epithelial stem cell lineage in live Drosophila ovarian tissue. We describe methods to generate transgenic flies expressing a pH biosensor, mCherry::pHluorin, image the biosensor using quantitative fluorescence imaging, generate standard curves, and convert fluorescence intensity values to pH values.

Changes in intracellular pH (pHi) play important roles in the regulation of many cellular functions, including metabolism, proliferation, and ...

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 ...

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with development, tissue homeostasis, and disease processes. Drosophila spermatocytes undergoing meiosis are ideal for this analysis because (1) whole Drosophila testes containing spermatocytes are relatively easy to prepare for microscopy, (2) the spermatocytes&#8217; large size makes them well suited for high resolution imaging, and (3) powerful Drosophila genetic tools can be integrated with in vivo analysis. Here, we present a readily accessible protocol for the preparation of whole testes from Drosophila third instar larvae and early pupae. We describe how to identify meiotic spermatocytes in prepared whole testes and how to image them live by time-lapse microscopy. Protocols for fixation and immunostaining whole testes are also provided. The use of larval testes has several advantages over available protocols that use adult testes for spermatocyte analysis. Most importantly, larval testes are smaller and less crowded with cells than adult testes, and this greatly facilitates high resolution imaging of spermatocytes. To demonstrate these advantages and the applications of the protocols, we present results showing the redistribution of the endoplasmic reticulum with respect to spindle microtubules during cell division in a single spermatocyte imaged by time-lapse confocal microscopy. The protocols can be combined with expression of any number of fluorescently tagged proteins or organelle markers, as well as gene mutations and other genetic tools, making this approach especially powerful for analysis of cell division mechanisms in the physiological context of whole tissues and organs.

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

The goal of this protocol is to analyze cell division in intact tissue by live and fixed cell microscopy using Drosophila meiotic spermatocytes. The protocol demonstrates how to isolate whole, intact testes from Drosophila larvae and early pupae, and how to process and mount them for microscopy.

Experimental analysis of cells dividing in living, intact tissues and organs is essential to our understanding of how cell division integrates with ...

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

The purpose of this protocol is to visualize intranuclear actin rods that assemble in live Drosophila melanogaster embryos following heat stress. Actin rods are a hallmark of a conserved, inducible Actin Stress Response (ASR) that accompanies human pathologies, including neurodegenerative disease. Previously, we showed that the ASR contributes to morphogenesis failures and reduced viability of developing embryos. This protocol allows the continued study of mechanisms underlying actin rod assembly and the ASR in a model system that is highly amenable to imaging, genetics and biochemistry. Embryos are collected and mounted on a coverslip to prepare them for injection. Rhodamine-conjugated globular actin (G-actinRed) is diluted and loaded into a microneedle. A single injection is made into the center of each embryo. After injection, embryos are incubated at elevated temperature and intranuclear actin rods are then visualized by confocal microscopy. Fluorescence recovery after photobleaching (FRAP) experiments may be performed on the actin rods; and other actin-rich structures in the cytoplasm can also be imaged. We find that G-actinRed polymerizes like endogenous G-actin and does not, on its own, interfere with normal embryo development. One limitation of this protocol is that care must be taken during injection to avoid serious injury to the embryo. However, with practice, injecting G-actinRed into Drosophila embryos is a fast and reliable way to visualize actin rods and can easily be used with flies of any genotype or with the introduction of other cellular stresses, including hypoxia and oxidative stress.

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos

The goal of this protocol is to inject Rhodamine-conjugated globular actin into Drosophila embryos and image intranuclear actin rod assembly following heat stress.

The purpose of this protocol is to visualize intranuclear actin rods that assemble in live Drosophila melanogaster embryos following heat stress. Actin ...

Live Imaging and Analysis of Muscle Contractions in Drosophila Embryo

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Live-imaging of the Drosophila Pupal Eye

Analysis of Zebrafish Larvae Skeletal Muscle Integrity with Evans Blue Dye

Observing Mitotic Division and Dynamics in a Live Zebrafish Embryo

Long-term Live Imaging of Drosophila Eye Disc

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

Live Imaging of Mouse Secondary Palate Fusion

Methods for Imaging Intracellular pH of the Follicle Stem Cell Lineage in Live Drosophila Ovarian Tissue

In Vivo Imaging of Muscle-tendon Morphogenesis in Drosophila Pupae

Preparation of Drosophila Larval and Pupal Testes for Analysis of Cell Division in Live, Intact Tissue

Imaging Intranuclear Actin Rods in Live Heat Stressed Drosophila Embryos