We gratefully acknowledge Eric Wieschaus, who provided the foundation on which this protocol was developed. Our work is supported by a Verna &amp; Marrs McLean Department of Biochemistry and Molecular Biology Start-up Award, Baylor College of Medicine.

1. Assemble Embryo Collection Cups
<ol>
<li>Cut the bottom off of a 100 mL Tri-corn beaker with a razor, making the edge as smooth as possible. The cups are easier to handle if you also trim the three corners off of the top, though this is not absolutely necessary.</li>
<li>Cut a square of wire mesh (6 cm x 6 cm). On a pre-warmed hot plate, inside of a fume hood, layer the square of wire mesh on top of a piece of heavy-duty aluminum foil. Push the cut bottom edge of the cup firmly onto the hot mesh. Wait for a few seconds, and lift the cup with the mesh now attached. If the foil also sticks, just peel it off.</li>
<li>Cool the cup overnight. Remove excess mesh with scissors, and sand off any sharp edges with fine-grain sandpaper.</li>
</ol>
2. Make Apple Juice Agar Plates
<ol>
<li>In a 6L flask, combine 100 g BD Bacto Agar and 3L distilled water. Autoclave the agar for 30 minutes on the slow exhaust setting.</li>
<li>In a 2L flask with a stir bar, combine 100 g sucrose, 1L apple juice, and 6 g p-Hydroxybenzoic acid. Heat the solution to boiling while stirring on a hot plate. Do not boil for more than 2 minutes. Allow the apple juice mixture to cool, and then add with stir bar to the agar. Mix completely.</li>
<li>Allow the combined solution to cool in a 60&deg;C water bath before pouring into 60x15 mm petri dishes. Alternatively, a peristaltic pump can be used to dispense the agar. Allow the plates to cool to room temperature for at least 4 hours. Stack in Rubbermaid containers with a layer of wet paper towels and store at 4&deg;C.</li>
</ol>
Notes:
<ul>
<li>In the fall or winter, it may be necessary to add an additional 100 mL of water to the agar.</li>
<li>The brand of petri dishes is important! Only the BD Falcon dishes fit the Tri-corn beakers. Specific ordering information can be found in the Materials table below.</li>
</ul>
3. Add GFP Flies to the Embryo Collection Cups
<ol>
<li>Expand GFP stocks according to your collection needs by setting up bottles of flies approximately two weeks prior to imaging. One bottle of flies is usually more than sufficient to fill one embryo collection cup with a minimum of 50 females and 30 males. For best laying, flies should be newly eclosed (i.e. less than 5 days post-hatching).</li>
<li>Make a yeast paste by filling a small cup with roughly equal parts Red Star active dry yeast and distilled water, and stir until the yeast is dissolved. The paste should approximate the consistency of wet peanut butter. More yeast or water can be added to alter the consistency. Yeast paste can be used for several days, and should be stored, covered, at 4&deg;C.</li>
<li>Once the yeast paste is ready, remove apple juice plates from 4&deg;C storage. Streak the apple juice plates with the yeast paste, and allow them to warm to 22-25&deg;C, which will encourage egg laying.</li>
<li>Fold a circular filter paper in half, then in half again. Trim to 8-9 cm diameter, and make accordion folds in the quarter circle. Unfold the circle, invert it, and insert it in a collection cup until it touches the mesh. Make sure that the paper is secure in the cup so that it does not fall and crush the flies. The filter paper is optional, but does provide a welcoming environment for mating, and maintains humidity in the cup.</li>
<li>Label a cup with stock name and date. Transfer the flies from the bottle to the collection cup by gently shaking the inverted bottle over the cup, and immediately cover the cup with a prepared apple juice plate. Secure the plate to the cup with a rubber band. Set cups mesh side up in an area with direct light. Make sure that no shadows fall on the cup, as flies will not lay well in the dark. </li>
<li>Change the apple juice plate as needed. Two-hour collections, at room temperature, work well for imaging cellularization.</li>
</ol>
4. Prepare a Mounting Chamber
<ol>
<li>To prepare a mounting chamber for the embryos, cut a piece of double-sided tape 2-3 cm long and place it on a slide, aligning the long axis of the tape and slide. Cut a second 2-3 cm long piece of double-sided tape and layer it on top of the other piece of tape, making sure their edges are aligned.</li>
<li>Using a razor blade, make two cuts approximately 3 mm apart in the center of the tape and perpendicular to the long axis of the slide. Remove the tape between the cuts to make a channel. Pipet a drop of Halocarbon 27 Oil into the channel. This channel is where the embryos will be mounted.</li>
</ol>
Notes:
<ul>
<li>Only the &frac12; inch Scotch double-sided tape is the right thickness to accommodate the embryos.</li>
<li>Halocarbon 27 Oil is oxygen permeable, and so allows oxygen exchange, while preventing embryo dehydration. Due to its refractive index, Halocarbon 27 Oil is also ideal for staging and imaging embryos.</li>
</ul>
5. Dechorionate Embryos
<ol>
<li>To collect and dechorionate embryos, pour enough 50% bleach on to the apple juice agar plate to fully submerge the entire surface. Using a dissecting microscope, such as the Zeiss Discovery V8 with transmitted light, watch for the embryos to release from their chorion. As soon as the chorion loosens and releases from a few embryos (30-60 seconds), pour bleach and embryos into a cell strainer.</li>
<li>Wash embryos in the strainer immediately and vigorously with distilled water from a squirt bottle. Dab strainer on paper towels. If any pink is seen on the towel after dabbing, continue washing the embryos with water.</li>
<li>Use a moist paintbrush to transfer 10-50 embryos to a clean apple juice plate (with no yeast paste). Wick away water with the torn edge of a paper towel, and immediately cover the embryos with a small amount of Halocarbon 27 Oil.</li>
</ol>
Notes:
<ul>
<li>Do not use Clorox bleach. Instead, use a brand such as Austin's A-1 Commercial Bleach, which is &lt;6% sodium hypochlorite.</li>
<li>Over-bleaching or insufficient rinsing will result in mushy embryos with irregular cellularization.</li>
</ul>
6. Stage and Mount Embryos
<ol>
<li>Using the dissecting microscope with transmitted light, follow the morphological guidelines of Bownes20 to stage embryos on the plate. Using forceps, transfer 5 mitotic cycle 11 or 12 embryos, corresponding to Bownes stage 4, to the channel of the mounting chamber. Arrange embryos in a line, with dorsal and ventral sides visible and lateral side down. Embryos are easily arranged in this orientation in oil alone. Do not crowd embryos together as they will deprive their neighbors of oxygen once the cover slip is applied below. (Leave approximately half of an embryo length between them).</li>
<li>Lay one edge of a 25x25 mm cover slip on the double-sided tape at one side of the channel. Drop the cover slip to cover the channel. If air is caught underneath, apply a small amount of Halocarbon 27 Oil at the edge of the cover slip. Capillary action will pull the oil into the channel and push out the air.</li>
</ol>
Notes:
<ul>
<li>Another excellent resource for staging embryos is the following: Campos-Ortega, J. A. and Hartenstein, V. (1985). The Embryonic development of Drosophila melanogaster. Springer-Verlag, Berlin. While this book is no longer published, used copies are regularly available on Amazon.com.</li>
<li>Always use the Halocarbon 27 Oil sparingly. This will make the mounting easier. It will also prevent accidental oozing of the oil onto the microscope objective in the following imaging steps.</li>
</ul>
7. Image Embryos
<ol>
<li>How you proceed here will of course be dictated by the cell shape change that you are imaging, and the question that you are addressing. For any shape change, place your mounting chamber on either an upright or inverted microscope, find your embryos by transmitted light, and then switch to confocal imaging to focus on a region of interest. Adhere to best practices for confocal microscopy, such as those discussed at Nikon's MicroscopyU (<a href="http://www.microscopyu.com/articles/confocal">http://www.microscopyu.com/articles/confocal</a>). Be particularly rigorous in exposing your living embryos to as little laser power as possible, to prevent photobleaching and phototoxicity.</li>
<li>For cellularization, we image on a Zeiss 710 laser scanning confocal microscope, using a C-Apochromat 40x/1.2W Corr objective. This microscope is housed in a temperature-controlled room, with temperatures ranging from 18-20&deg;C. While temperature control is not absolutely necessary, it is better for the microscope and makes the imaging more consistent. We regularly image in a single plane, near the middle of the embryo, to follow the plasma membrane invaginations in cross-section (Figure 2). For simply following invagination dynamics, 30-60 second intervals are sufficient for time-lapse imaging, and embryos should withstand imaging for >90 minutes with no obvious signs of oxygen deprivation.</li>
<li>After acquiring the images, they can be analyzed using any number of image analysis packages, including freeware from the National Institutes of Health, called ImageJ (<a href="http://rsbweb.nih.gov/ij">http://rsbweb.nih.gov/ij</a>). The data can then be presented as movies (Movie 1), sequences (Figure 2), or kymographs21.</li>
</ol>
Note:
<ul>
<li>The water immersion C-Apochromat 40x/1.2W Corr objective is well-suited for imaging near the embryo mid-section due to its high aperture, which limits aberrations for deep tissue imaging in aqueous samples, and gives high resolution and high fluorescent signal. In addition, this objective has a very long working distance (220 &mu;m). However, other water or glycerine immersion objectives will perform well, particularly if the cell shape change of interest can be imaged near the embryo surface.</li>
</ul>
8. Alternative Method: Mount Embryos with "embryo-glue"
<ol>
<li>To image a cell shape change other than cellularization, it may be necessary to mount embryos in a specific orientation that is not easily maintained in oil alone. To do so, use an alternate mounting method to the one described previously. Start by making "embryo-glue" by combining 20 cm of double-sided tape with 250 &mu;L of heptane in a scintillation vial. Place the scintillation vial on a nutator or rotating platform, and mix overnight.</li>
<li>Dip a yellow pipette tip into the embryo glue, and then trace the tip along a slide, leaving a trail of glue. While the heptane is evaporating, align your staged embryos in the required orientation on a block of agar. (Remember that the embryo surface to be imaged should be facing the agar. For example, to image cell shape change during ventral furrow formation, mount embryos with their ventral side facing the agar.)</li>
<li>Invert the slide with glue, and gently press the trail of glue against the embryos on the agar block. Now invert the slide again. The embryos will be stuck in the appropriate orientation. Add two layers of double-sided tape on either side of the embryos, cover them with Halocarbon 27 Oil, and apply a coverslip. Proceed with the imaging as described above.</li>
</ol>
9. Representative Results:
If the embryos are healthy and the imaging is optimal, then cellularization should take 50-60 minutes, and the plasma membrane invaginations should ingress almost 40 microns. However, if the embryos are over-bleached, oxygen deprived or damaged by phototoxicity, then invagination will either slow or stop, particularly in the imaged area. Such deterioration of embryo health often results in altered development, and a failure to hatch as larvae. Thus, for a rigorous test to assay embryo health after imaging, keep your slides in a humidified chamber and watch for hatching the next day.
<img src="/files/ftp_upload/2503/2503fig1.jpg" alt="Figure 1" /> 
Figure 1. Workflow from embryo collection to imaging. The workflow of the protocol can be broken down into four main phases. In the first phase, all the individual supplies and components are prepared, and then the embryo collection cup, apple juice agar plate and GFP flies are put together to create the embryo-laying environment. In the second phase, the eggs and embryos that are laid on the apple juice agar plates are collected. In the third phase, the embryos are removed from the plate, staged and transferred to the mounting chamber. In the fourth phase, the mounted embryos are imaged on a confocal microscope.
<img src="/files/ftp_upload/2503/2503fig2.jpg" alt="Figure 2" /> 
Figure 2. Representative data from time-lapse imaging of cellularization. Embryos are mounted with dorsal (D) and ventral (V) sides clearly visible, and are imaged near their middle to follow the plasma membrane invaginations in cross-section. The embryo shown here expresses a GFP-Myosin-2 probe6, which concentrates at the tips of the plasma membrane invaginations. Thus, tracking the ingression of this front over time gives the rate at which the plasma membrane invaginates. The 0:00 minute time point corresponds to cellularization onset. Shortly after the 56:00 minute time point, gastrulation starts on the ventral side of the embryo. Bar is 40 microns.
Movie 1. Representative movie from time-lapse imaging of cellularization. This movie corresponds to Figure 2. To record the entire process of cellularization, imaging started in the prior mitotic cycle 13, capturing pseudocleavage furrow regression, and continued until gastrulation movements were seen on the ventral side of the embryo. Images were collected at one minute intervals. The intensities were increased post-acquisition to make it easier to see the gastrulation movements. 
<a href="https://www.jove.com/files/ftp_upload/2503/Movie%201.mov">Click here for movie</a>
<table border="1">
<tbody>
 <tr>
 <td>Developmental event and timing*</td>
 <td>Cell shape changes related to</td>
 <td>A link to disease or human health</td>
 <td>Recent references with live imaging</td>
 </tr>
 <tr>
 <td>Pseudo-cleavage furrow formation 
 (4; 90 minutes pf)</td>
 <td>Cytokinesis</td>
 <td>Polyploidy and cancer progression1</td>
 <td>Mavrakis et al., 20098 
 Cao et al., 20109</td>
 </tr>
 <tr>
 <td>Cellularization 
 (5; 130 minutes pf)</td>
 <td>Cytokinesis</td>
 <td>Polyploidy and cancer progression1</td>
 <td>Cao et al., 200810 
 Sokac &amp; Wieschaus, 200811</td>
 </tr>
 <tr>
 <td>Ventral furrow formation; Mesoderm invagination 
 (6; 180 minutes pf)</td>
 <td>Apical constriction; Epithelial-mesenchymal transition</td>
 <td>Cancer metastasis2</td>
 <td>Fox &amp; Peifer, 200712 
 Martin et al., 200913</td>
 </tr>
 <tr>
 <td>Germband extension 
 (7; 195 minutes pf)</td>
 <td>Convergent extension</td>
 <td>Neural tube defects3</td>
 <td>Bertet et al., 200414 
 Blankenship et al., 200615</td>
 </tr>
 <tr>
 <td>Tracheogenesis 
 (11; 320 minutes pf)</td>
 <td>Epithelial tube formation and branching</td>
 <td>Angiogenesis4</td>
 <td>Caussinus et al., 200816 
 Gervais &amp; Casanova, 201017</td>
 </tr>
 <tr>
 <td>Dorsal closure 
 (14; 620 minutes pf)</td>
 <td>Apical constriction</td>
 <td>Wound healing5</td>
 <td>Gorfinkiel et al., 200918 
 Solon et al., 200919</td>
 </tr>
</tbody>
</table>
Table 1. Examples of cell shape changes imaged in living fly embryos 
*The Bownes stage number and time post-fertilization (pf), when each event starts, are listed according to Campos-Ortega,1985. 

<table border="1">
<tbody>
 <tr>
 <td>Fly stock</td>
 <td>Labels</td>
 <td>Original reference </td>
 </tr>
 <tr>
 <td>Spider-GFP (95-1)</td>
 <td>Plasma membrane</td>
 <td>Morin et al., 200122</td>
 </tr>
 <tr>
 <td>Resille-GFP (117-2)</td>
 <td>Plasma membrane</td>
 <td>Morin et al., 200122</td>
 </tr>
 <tr>
 <td>GAP43-Venus</td>
 <td>Plasma membrane</td>
 <td>Mavrakis et al., 20098</td>
 </tr>
 <tr>
 <td>Spaghetti Squash-GFP (Sqh-GFP)</td>
 <td>Myosin-2</td>
 <td>Royou et al., 20026</td>
 </tr>
 <tr>
 <td>E-cadherin-GFP (Ecad-GFP)</td>
 <td>Cell-cell junctions</td>
 <td>Oda et al., 200123</td>
 </tr>
 <tr>
 <td>GFP-Moesin</td>
 <td>F-actin</td>
 <td>Kiehart et al., 200024</td>
 </tr>
 <tr>
 <td>Utrophin-Venus (Utro-Venus)</td>
 <td>F-actin</td>
 <td>Sokac et al., unpublished results</td>
 </tr>
</tbody>
</table>
Table 2. Useful stocks for imaging cell shape change in fly embryos

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No conflicts of interest declared.

The protocol described herein will permit the live, confocal imaging of a number of cell shape changes in the developing fly embryo. GFP stocks for imaging can be prepared by an individual lab (Table 2), but many such stocks are also publicly available from centers such as Bloomington Drosophila Stock Center at Indiana University (<a href="http://flystocks.bio.indiana.edu">http://flystocks.bio.indiana.edu</a>) and FlyTrap Stock Center at Yale University (<a href="http://flytrap.med.yale.edu">http://flytrap.med....

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imaging cell shape change in living drosophila embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging. Cell shape changes in the fly are analogous to those in higher organisms, and they drive tissue morphogenesis. So, in many cases, their study has direct implications for understanding human disease (Table 1)1-5. On the sub-cellular scale, these cell shape changes are the product of activities ranging from gene expression to signal transduction, cell polarity, cytoskeletal remodeling and membrane trafficking. Thus, the Drosophila embryo provides not only the context to evaluate cell shape changes as they relate to tissue morphogenesis, but also offers a completely physiological environment to study the sub-cellular activities that shape cells.
The protocol described here is designed to image a specific cell shape change called cellularization. Cellularization is a process of dramatic plasma membrane growth, and it ultimately converts the syncytial embryo into the cellular blastoderm. That is, at interphase of mitotic cycle 14, the plasma membrane simultaneously invaginates around each of ~6000 cortically anchored nuclei to generate a sheet of primary epithelial cells. Counter to previous suggestions, cellularization is not driven by Myosin-2 contractility6, but is instead fueled largely by exocytosis of membrane from internal stores7. Thus, cellularization is an excellent system for studying membrane trafficking during cell shape changes that require plasma membrane invagination or expansion, such as cytokinesis or transverse-tubule (T-tubule) morphogenesis in muscle.
Note that this protocol is easily applied to the imaging of other cell shape changes in the fly embryo, and only requires slight adaptations such as changing the stage of embryo collection, or using "embryo glue" to mount the embryo in a specific orientation (Table 1)8-19. In all cases, the workflow is basically the same (Figure 1). Standard methods for cloning and Drosophila transgenesis are used to prepare stable fly stocks that express a protein of interest, fused to Green Fluorescent Protein (GFP) or its variants, and these flies provide a renewable source of embryos. Alternatively, fluorescent proteins/probes are directly introduced into fly embryos via straightforward micro-injection techniques9-10. Then, depending on the developmental event and cell shape change to be imaged, embryos are collected and staged by morphology on a dissecting microscope, and finally positioned and mounted for time-lapse imaging on a confocal microscope.

Watch this Scientific Journal Video about Imaging Cell Shape Change in Living Drosophila Embryos at JoVE.com

Imaging Cell Shape Change in Living Drosophila Embryos

Early development of the fruit fly, Drosophila melanogaster, is characterized by a number of cell shape changes that are well suited for imaging approaches. This article will describe basic tools and methods required for live confocal imaging of Drosophila embryos, and will focus on a cell shape change called cellularization.

Imaging Cell Shape Change in Living Drosophila Embryos

The developing Drosophila melanogaster embryo undergoes a number of cell shape changes that are highly amenable to live confocal imaging.  Cell shape ...

imaging-cell-shape-change-in-living-drosophila-embryos

Research

JoVE Journal

Biology

14.2K 次观看.  Baylor College of Medicine (BCM). 果蝇，果蝇，早期发展的特点是由许多细胞形状的变化，以及成像方法适合。本文将描述活果蝇胚胎的共焦成像所需的基本工具和方法，将集中在细胞形态变化称为cellularization。

视频: Imaging Cell Shape Change in Living Drosophila Embryos

Preparation of Neuronal Cultures from Midgastrula Stage Drosophila Embryos

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos and their dechorionation using bleach are demonstrated. Using a glass pipet attached to a mouth suction tube, we illustrate the removal of all cells from single embryos. The method for dispersing cells from each embyro into a small (5 l) drop of medium on an uncoated glass coverslip is demonstrated. A view through the microscope at 1 hour after plating illustrates the preferred cell density. Most of the cells that survive when grown in defined medium are neuroblasts that divide one or more times in culture before extending neuritic processes by 12-24 hours. A view through the microscope illustrates the level of neurite outgrowth and branching expected in a healthy culture at 2 days in vitro. The cultures are grown in a simple bicarbonate based defined medium, in a 5% CO2 incubator at 22-24&deg;C. Neuritic processes continue to elaborate over the first week in culture and when they make contact with neurites from neighboring cells they often form functional synaptic connections. Neurons in these cultures express voltage-gated sodium, calcium, and potassium channels and are electrically excitable. This culture system is useful for studying molecular genetic and environmental factors that regulate neuronal differentiation, excitability, and synapse formation/function.

This video demonstrates the preparation of primary neuronal cultures from midgastrula stage Drosophila embryos. Views of live cultures show cells 1 hour after plating and differentiated neurons after 2 days of growth in a bicarbonate-based defined medium. The neurons are electrically excitable and form synaptic connections.

This video illustrates the procedure for making primary neuronal cultures from midgastrula stage Drosophila embryos. The methods for collecting embryos ...

Two-photon axotomy and time-lapse confocal imaging in live zebrafish embryos

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo. Here we describe a method to mount zebrafish embryos for long-term imaging and demonstrate how to automate the capture of time-lapse images using a confocal microscope. We also describe a method to create controlled, precise damage to individual branches of peripheral sensory axons in zebrafish using the focused power of a femtosecond laser mounted on a two-photon microscope. The parameters for successful two-photon axotomy must be optimized for each microscope. We will demonstrate two-photon axotomy on both a custom built two-photon microscope and a Zeiss 510 confocal/two-photon to provide two examples.

Zebrafish trigeminal sensory neurons can be visualized in a transgenic line expressing GFP driven by a sensory neuron specific promoter 1. We have adapted this zebrafish trigeminal model to directly observe sensory axon regeneration in living zebrafish embryos. Embryos are anesthetized with tricaine and positioned within a drop of agarose as it solidifies. Immobilized embryos are sealed within an imaging chamber filled with phenylthiourea (PTU) Ringers. We have found that embryos can be continuously imaged in these chambers for 12-48 hours. A single confocal image is then captured to determine the desired site of axotomy. The region of interest is located on the two-photon microscope by imaging the sensory axons under low, non-damaging power. After zooming in on the desired site of axotomy, the power is increased and a single scan of that defined region is sufficient to sever the axon. Multiple location time-lapse imaging is then set up on a confocal microscope to directly observe axonal recovery from injury.

Here we describe a method for mounting zebrafish embryos for long-term imaging, two-photon imaging and tissue-damage techniques, and time-lapse confocal imaging.

Zebrafish have long been utilized to study the cellular and molecular mechanisms of development by time-lapse imaging of the living transparent embryo.  ...

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as tissue morphogenesis, cell-cell adhesion, or cell death. Drosophila embryos expressing GFP are readily imaged using either stereoscopic or confocal microscopy. A goal of any live-imaging protocol is to minimize detrimental effects such as dehydration and hypoxia. Previous protocols for preparing Drosophila embryos for live-imaging analysis have involved placing dechorionated embryos in halocarbon oil and sandwiching them between a halocarbon gas-permeable membrane and a coverslip1-3. The introduction of compression through mounting embryos in this manner represents an undesirable complication for any biomechanical-based analysis of morphogenesis. Our method, which we call the hanging drop protocol, results in excellent viability of embryos during live imaging and does not require that embryos be compressed. Briefly, the hanging drop protocol involves the placement of embryos in a drop of halocarbon oil that is suspended from a coverslip, which is, in turn, fixed in position over a humid chamber. In addition to providing gas exchange and preventing dehydration, this arrangement takes advantage of the buoyancy of embryos in halocarbon oil to prevent them from drifting out of position during timelapse acquisition. This video describes in detail how to collect and prepare Drosophila embryos for live imaging using the hanging drop protocol. This protocol is suitable for imaging dechorionated embryos using stereomicroscopy or any upright compound fluorescence microscope.

The Preparation of Drosophila Embryos for Live-Imaging Using the Hanging Drop Protocol

A simple, inexpensive, and effective method of preparing Drosophila embryos for live-imaging analysis is presented. Our protocol provides humidity and gas exchange and does not compress the Drosophila embryo. This method is suitable for GFP-based live imaging of Drosophila embryos using a stereomicroscope or upright compound microscope.

Green fluorescent protein (GFP)-based timelapse live-imaging is a powerful technique for studying the genetic regulation of dynamic processes such as ...

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

The morphogenesis of the Drosophila embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell shape changes during embryonic development. One of the challenges faced in studying this structure is that the lumen of the heart tube, as well as the membrane features that are crucial to heart tube formation, are difficult to visualize in whole mount embryos, due to the small size of the heart tube and intra-lumenal space relative to the embryo. The use of transmission electron microscopy allows for higher magnification of these structures and gives the advantage of examining the embryos in cross section, which easily reveals the size and shape of the lumen. In this video, we detail the process for reliable fixation, embedding, and sectioning of late stage Drosophila embryos in order to visualize the heart tube lumen as well as important cellular structures including cell-cell junctions and the basement membrane.

Preparation of embryos for Electron Microscopy of the Drosophila embryonic heart tube

We describe a process for fixation, embedding, sectioning, and imaging of late stage Drosophila embryos for Trasmission Electron Microscopy of the embryonic heart tube. This technique allows for the visualization of the heart tube lumen as well as the basement membrane, which lines the lumen of the heart.

The morphogenesis of the Drosophila  embryonic heart tube has emerged as a valuable model system for studying cell migration, cell-cell adhesion and cell ...

In-vivo Centrifugation of Drosophila Embryos

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in buoyant density. However, when cells are disrupted prior to centrifugation, proteins and organelles in this non-native environment often inappropriately stick to each other. Here we describe a method to separate organelles by density in intact, living Drosophila embryos. Early embryos before cellularization are harvested from population cages, and their outer egg shells are removed by treatment with 50% bleach. Embryos are then transferred to a small agar plate and inserted, posterior end first, into small vertical holes in the agar. The plates containing embedded embryos are centrifuged for 30 min at 3000g. The agar supports the embryos and keeps them in a defined orientation. Afterwards, the embryos are dug out of the agar with a blunt needle. Centrifugation separates major organelles into distinct layers, a stratification easily visible by bright-field microscopy. A number of fluorescent markers are available to confirm successful stratification in living embryos. Proteins associated with certain organelles will be enriched in a particular layer, demonstrating colocalization. Individual layers can be recovered for biochemical analysis or transplantation into donor eggs. This technique is applicable for organelle separation in other large cells, including the eggs and oocytes of diverse species.

In-vivo Centrifugation of Drosophila Embryos

We describe a method to separate organelles by density in living Drosophila embryos. Embryos are embedded in agar and centrifuged. This technique yields reproducible separation of major organelles along the anterior-posterior embryo axis. This method facilitates colocalization experiments and yields organelle fractions for biochemical analysis and transplantation experiments.

A major strategy for purifying and isolating different types of intracellular organelles is to separate them from each other based on differences in ...

Upright Imaging of Drosophila Embryos

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current techniques used to view such processes are somewhat limited. The following protocol describes a new technique for mounting fixed and labeled Drosophila embryos for coronal viewing with confocal imaging. This method consists of embedding embryos between two layers of glycerin jelly mounting media, and imaging jelly strips positioned upright. The first step for sandwiching the embryos is to make a thin bedding of glycerin jelly on a slide. Next, embryos are carefully aligned on this surface and covered with a second layer of jelly. After the second layer is solidified, strips of jelly are cut and flipped upright for imaging. Alternatives are described for visualizing the embryos depending upon the type of microscope stand to be used. Since all cells along the dorsal-ventral axis are imaged within a single confocal Z-plane, our method allows precise measurement and comparison of fluorescent signals without photobleaching or light scattering common to 3D reconstructions of longitudinally mounted embryos.

Upright Imaging of Drosophila Embryos

Here we present a mounting protocol for stained Drosophila embryos in an upright position that allows imaging of cross-sections using Confocal microscopy.

Several well-known morphogenetic gradients and cellular movements occur along the dorsal/ventral axis of the Drosophila embryo. However, the current ...

Primary Cell Cultures from Drosophila Gastrula Embryos

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos. In brief, a large amount of staged embryos from young and healthy flies are collected, sterilized, and then physically dissociated into a single cell suspension using a glass homogenizer. After being plated on culture plates or chamber slides at an appropriate density in culture medium, these cells can further differentiate into several morphologically-distinct cell types, which can be identified by their specific cell markers. Furthermore, we present conditions for treating these cells with double stranded (ds) RNAs to elicit gene knockdown. Efficient RNAi in Drosophila primary cells is accomplished by simply bathing the cells in dsRNA-containing culture medium. The ability to carry out effective RNAi perturbation, together with other molecular, biochemical, cell imaging analyses, will allow a variety of questions to be answered in Drosophila primary cells, especially those related to differentiated muscle and neuronal cells.

Primary Cell Cultures from Drosophila Gastrula Embryos

We provide a detailed protocol for preparing primary cells dissociated from Drosophila embryos. The ability to carry out the effective RNAi perturbation, together with other molecular, biochemical and cell imaging methods will allow a variety of questions to be addressed in Drosophila primary cells.

Here we describe a method for preparing and culturing primary cells dissociated from Drosophila gastrula embryos.   In brief, a large amount of staged ...

Isolation and Purification of Kinesin from Drosophila Embryos

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a variety of neurodegenerative diseases, understanding microtubule based motor transport and its regulation will likely ultimately lead to improved therapeutic approaches. Kinesin-1 is a eukaryotic motor protein which moves in an anterograde (plus-end) direction along microtubules (MTs), powered by ATP hydrolysis. Here we report a detailed purification protocol to isolate active full length kinesin from Drosophila embryos, thus allowing the combination of Drosophila genetics with single-molecule biophysical studies. Starting with approximately 50 laying cups, with approximately 1000 females per cup, we carried out overnight collections. This provided approximately 10 ml of packed embryos. The embryos were bleach dechorionated (yielding approximately 9 grams of embryos), and then homogenized. After disruption, the homogenate was clarified using a low speed spin followed by a high speed centrifugation. The clarified supernatant was treated with GTP and taxol to polymerize MTs. Kinesin was immobilized on polymerized MTs by adding the ATP analog, 5'-adenylyl imidodiphosphate at room temperature. After kinesin binding, microtubules were sedimented via high speed centrifugation through a sucrose cushion. The microtubule pellet was then re-suspended, and this process was repeated. Finally, ATP was added to release the kinesin from the MTs. High speed centrifugation then spun down the MTs, leaving the kinesin in the supernatant. This kinesin was subjected to a centrifugal filtration using a 100 KD cut off filter for further purification, aliquoted, snap frozen in liquid nitrogen, and stored at -80 &deg;C. SDS gel electrophoresis and western blotting was performed using the purified sample. The motor activity of purified samples before and after the final centrifugal filtration step was evaluated using an in vitro single molecule microtubule assay. The kinesin fractions before and after the centrifugal filtration showed processivity as previously reported in literature. Further experiments are underway to evaluate the interaction between kinesin and other transport related proteins.

Isolation and Purification of Kinesin from Drosophila Embryos

This is a protocol to isolate active full length Kinesin from Drosophila embryos for single-molecule biophysical studies. We show how to collect embryos, make the embryo lysate, and then polymerize microtubules (MTs). Kinesin is purified by immobilizing it on the MTs, spinning down the Kinesin-MT complexes, and then releasing the kinesin from the MTs via ATP addition.

Motor proteins move cargos along microtubules, and transport them to specific sub-cellular locations. Because altered transport is suggested to underlie a ...

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle microtubules to prevent anaphase onset until the chromosomes are properly connected. Cells use this mechanism to prevent aneuploidy or genomic instability, and hence cancers and other human diseases like birth defects and Alzheimer's1. A number of the SAC components such as Mad1, Mad2, Bub1, BubR1, Bub3, Mps1, Zw10, Rod and Aurora B kinase have been identified and they are all kinetochore dynamic proteins2. Evidence suggests that the kinetochore is where the SAC signal is initiated. The SAC prime regulatory target is Cdc20. Cdc20 is one of the essential APC/C (Anaphase Promoting Complex or Cyclosome) activators3 and is also a kinetochore dynamic protein4-6. When activated, the SAC inhibits the activity of the APC/C to prevent the destruction of two key substrates, cyclin B and securin, thereby preventing the metaphase to anaphase transition7,8. Exactly how the SAC signal is initiated and assembled on the kinetochores and relayed onto the APC/C to inhibit its function still remains elusive.
Drosophila is an extremely tractable experimental system; a much simpler and better-understood organism compared to the human but one that shares fundamental processes in common. It is, perhaps, one of the best organisms to use for bio-imaging studies in living cells, especially for visualization of the mitotic events in space and time, as the early embryo goes through 13 rapid nuclear division cycles synchronously (8-10 minutes for each cycle at 25 &deg;C) and gradually organizes the nuclei in a single monolayer just underneath the cortex9.
Here, I present a bio-imaging method using transgenic Drosophila expressing GFP (Green Fluorescent Protein) or its variant-targeted proteins of interest and a Leica TCS SP2 confocal laser scanning microscope system to study the SAC function in flies, by showing images of GFP fusion proteins of some of the SAC components, Cdc20 and Mad2, as the example.

Studying Mitotic Checkpoint by Illustrating Dynamic Kinetochore Protein Behavior and Chromosome Motion in Living Drosophila Syncytial Embryos

The kinetochore is where the SAC initiates its signal monitoring the mitotic segregation of the sister chromatids. A method is described to visualize the recruitment and turnover of one of the kinetochore proteins and its coordination with the chromosome motion in Drosophila embryos using a Leica laser scanning confocal system.

The spindle assembly checkpoint (SAC) mechanism is an active signal, which monitors the interaction between chromosome kinetochores and spindle ...

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

The proper elimination of unwanted or aberrant cells through apoptosis and subsequent phagocytosis (apoptotic cell clearance) is crucial for normal development in all metazoan organisms. Apoptotic cell clearance is a highly dynamic process intimately associated with cell death; unengulfed apoptotic cells are barely seen in vivo under normal conditions. In order to understand the different steps of apoptotic cell clearance and to compare 'professional' phagocytes - macrophages and dendritic cells to 'non-professional' - tissue-resident neighboring cells, in vivo live imaging of the process is extremely valuable. Here we describe a protocol for studying apoptotic cell clearance in live Drosophila embryos. To follow the dynamics of different steps in phagocytosis we use specific markers for apoptotic cells and phagocytes. In addition, we can monitor two phagocyte systems in parallel: 'professional' macrophages and 'semi-professional' glia in the developing central nervous system (CNS). The method described here employs the Drosophila embryo as an excellent model for real time studies of apoptotic cell clearance.

Live Imaging of Apoptotic Cell Clearance during Drosophila Embryogenesis

Here we describe an effective method for studying dynamics of apoptotic cell clearance in vivo. This method employs live Drosophila embryos as a powerful model for monitoring phagocytosis of apoptotic cells using specific labeling of apoptotic cells and phagocytes.

The  proper elimination of unwanted or aberrant cells through apoptosis and  subsequent phagocytosis (apoptotic cell clearance) is crucial for normal  ...