Name: imaging cell shape change in living drosophila embryos
Uploaded: 2011-03-30T13:00:00
Description: Watch this Scientific Journal Video about Imaging Cell Shape Change in Living Drosophila Embryos at JoVE.com

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 f...

<ol>
	<li>Sagona, A. P., Stenmark, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytokinesis+and+cancer.">Cytokinesis and cancer.</a> FEBS Lett. 584, 2652-2661 (2010).</li><li>Baum, B., Settleman, J., Quinlan, M. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transitions+between+epithelial+and+mesenchymal+states+in+development+and+disease.">Transitions between epithelial and mesenchymal states in development and disease.</a> Semin Cell Dev Biol. 19, 294-308 (2008).</li><li>Kibar, Z., Capra, V., Gros, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Toward+understanding+the+genetic+basis+of+neural+tube+defects.">Toward understanding the genetic basis of neural tube defects.</a> Clin Genet. 71, 295-310 (2007).</li><li>Jazwinska, A., Ribeiro, C., Affolter, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Epithelial+tube+morphogenesis+during+Drosophila+tracheal+development+requires+Piopio,+a+luminal+ZP+protein.">Epithelial tube morphogenesis during Drosophila tracheal development requires Piopio, a luminal ZP protein.</a> Nat Cell Biol. 5, 895-901 (2003).</li><li>Martin, P., Parkhurst, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Parallels+between+tissue+repair+and+embryo+morphogenesis.">Parallels between tissue repair and embryo morphogenesis.</a> Development. 131, 3021-3034 (2004).</li><li>Royou, A., Field, C., Sisson, J. C., Sullivan, W., Karess, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Reassessing+the+role+and+dynamics+of+nonmuscle+myosin+II+during+furrow+formation+in+early+Drosophila+embryos.">Reassessing the role and dynamics of nonmuscle myosin II during furrow formation in early Drosophila embryos.</a> Mol Biol Cell. 15, 838-850 (2004).</li><li>Lecuit, T., Wieschaus, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Polarized+insertion+of+new+membrane+from+a+cytoplasmic+reservoir+during+cleavage+of+the+Drosophila+embryo.">Polarized insertion of new membrane from a cytoplasmic reservoir during cleavage of the Drosophila embryo.</a> J Cell Biol. 150, 849-860 (2000).</li><li>Mavrakis, M., Rikhy, R., Lippincott-Schwartz, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Plasma+membrane+polarity+and+compartmentalization+are+established+before+cellularization+in+the+fly+embryo.">Plasma membrane polarity and compartmentalization are established before cellularization in the fly embryo.</a> Dev Cell. 16, 93-104 (2009).</li><li>Cao, J., Crest, J., Fasulo, B., Sullivan, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cortical+Actin+Dynamics+Facilitate+Early-Stage+Centrosome+Separation.">Cortical Actin Dynamics Facilitate Early-Stage Centrosome Separation.</a> Curr Biol. , (2010).</li><li>Cao, J., Albertson, R., Riggs, B., Field, C. M., Sullivan, W. N. u. f. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=a+Rab11+effector,+maintains+cytokinetic+furrow+integrity+by+promoting+local+actin+polymerization.">a Rab11 effector, maintains cytokinetic furrow integrity by promoting local actin polymerization.</a> J Cell Biol. 182, 301-313 (2008).</li><li>Sokac, A. M., Wieschaus, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Local+actin-dependent+endocytosis+is+zygotically+controlled+to+initiate+Drosophila+cellularization.">Local actin-dependent endocytosis is zygotically controlled to initiate Drosophila cellularization.</a> Dev Cell. 14, 775-786 (2008).</li><li>Fox, D. T., Peifer, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Abelson+kinase+(Abl)+and+RhoGEF2+regulate+actin+organization+during+cell+constriction+in+Drosophila.">Abelson kinase (Abl) and RhoGEF2 regulate actin organization during cell constriction in Drosophila.</a> Development. 134, 567-578 (2007).</li><li>Martin, A. C., Kaschube, M., Wieschaus, E. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pulsed+contractions+of+an+actin-myosin+network+drive+apical+constriction.">Pulsed contractions of an actin-myosin network drive apical constriction.</a> Nature. 457, 495-499 (2009).</li><li>Bertet, C., Sulak, L., Lecuit, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Myosin-dependent+junction+remodelling+controls+planar+cell+intercalation+and+axis+elongation.">Myosin-dependent junction remodelling controls planar cell intercalation and axis elongation.</a> Nature. 429, 667-671 (2004).</li><li>Blankenship, J. T., Backovic, S. T., Sanny, J. S., Weitz, O., Zallen, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multicellular+rosette+formation+links+planar+cell+polarity+to+tissue+morphogenesis.">Multicellular rosette formation links planar cell polarity to tissue morphogenesis.</a> Dev Cell. 11, 459-470 (2006).</li><li>Caussinus, E., Colombelli, J., Affolter, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tip-cell+migration+controls+stalk-cell+intercalation+during+Drosophila+tracheal+tube+elongation.">Tip-cell migration controls stalk-cell intercalation during Drosophila tracheal tube elongation.</a> Curr Biol. 18, 1727-1734 (2008).</li><li>Gervais, L., Casanova, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vivo+coupling+of+cell+elongation+and+lumen+formation+in+a+single+cell.">In vivo coupling of cell elongation and lumen formation in a single cell.</a> Curr Biol. 20, 359-366 (2010).</li><li>Gorfinkiel, N., Blanchard, G. B., Adams, R. J., Arias, M. a. r. t. i. n. e. z., A, . <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+control+of+global+cell+behaviour+during+dorsal+closure+in+Drosophila.">Mechanical control of global cell behaviour during dorsal closure in Drosophila.</a> Development. 136, 1889-1898 (2009).</li><li>Solon, J., Kaya-Copur, A., Colombelli, J., Brunner, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pulsed+forces+timed+by+a+ratchet-like+mechanism+drive+directed+tissue+movement+during+dorsal+closure.">Pulsed forces timed by a ratchet-like mechanism drive directed tissue movement during dorsal closure.</a> Cell. 137, 1331-1342 (2009).</li><li>Bownes, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+photographic+study+of+development+in+the+living+embryo+of+Drosophila+melanogaster.">A photographic study of development in the living embryo of Drosophila melanogaster.</a> J Embryol Exp Morphol. 33, 789-801 (1975).</li><li>Sokac, A. M., Wieschaus, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Zygotically+controlled+F-actin+establishes+cortical+compartments+to+stabilize+furrows+during+Drosophila+cellularization.">Zygotically controlled F-actin establishes cortical compartments to stabilize furrows during Drosophila cellularization.</a> J Cell Sci. 121, 1815-1824 (2008).</li><li>Morin, X., Daneman, R., Zavortink, M., Chia, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+protein+trap+strategy+to+detect+GFP-tagged+proteins+expressed+from+their+endogenous+loci+in+Drosophila.">A protein trap strategy to detect GFP-tagged proteins expressed from their endogenous loci in Drosophila.</a> Proc Natl Acad Sci U S A. 98, 15050-15055 (2001).</li><li>Oda, H., Tsukita, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Real-time+imaging+of+cell-cell+adherens+junctions+reveals+that+Drosophila+mesoderm+invagination+begins+with+two+phases+of+apical+constriction+of+cells.">Real-time imaging of cell-cell adherens junctions reveals that Drosophila mesoderm invagination begins with two phases of apical constriction of cells.</a> J Cell Sci. 114, 493-501 (2001).</li><li>Kiehart, D. P., Galbraith, C. G., Edwards, K. A., Rickoll, W. L., Montague, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Multiple+forces+contribute+to+cell+sheet+morphogenesis+for+dorsal+closure+in+Drosophila.">Multiple forces contribute to cell sheet morphogenesis for dorsal closure in Drosophila.</a> J Cell Biol. 149, 471-490 (2000).</li></ol>

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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		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Slides</td>
<td>&nbsp;</td>
<td>Fisherbrand/Fisher Scientific</td>
<td>12-550-343</td>
<td>&nbsp;</td>
<tr>
<td>Cover slips 25x25</td>
<td>&nbsp;</td>
<td>Fisher Scientific (Corning #2865-25)</td>
<td>1 2-524C</td>
<td>&nbsp;</td>
<tr>
<td>Squirt bottles (H2O)</td>
<td>&nbsp;</td>
<td>Fisher Scientific</td>
<td>02-897-11</td>
<td>&nbsp;</td>
<tr>
<td>50 ml Falcon tubes</td>
<td>&nbsp;</td>
<td>Fisher Scientific (BD# 352070)</td>
<td>14-432-22</td>
<td>&nbsp;</td>
<tr>
<td>Bulbs for small pipets, 1 mL</td>
<td>&nbsp;</td>
<td>Fisherbrand/Fisher Scientific</td>
<td>03-448-21</td>
<td>&nbsp;</td>
<tr>
<td>Scintillation vials with caps</td>
<td>&nbsp;</td>
<td>VWR (Wheaton #986546)</td>
<td>66021-533</td>
<td>&nbsp;</td>
<tr>
<td>Tri-Corn Beakers, 100 mL</td>
<td>&nbsp;</td>
<td>Electron Microscopy Sciences</td>
<td>60970</td>
<td>&nbsp;</td>
<tr>
<td>BD Falcon Petri dish 60x15mm </td>
<td>&nbsp;</td>
<td>Fisher Scientific (BD# 351007)</td>
<td>08757 100B</td>
<td>&nbsp;</td>
<tr>
<td>BD Falcon Cell strainer</td>
<td>&nbsp;</td>
<td>Fisher Scientific (BD#352350)</td>
<td>08-771-2</td>
<td>&nbsp;</td>
<tr>
<td>Yellow pipet tips</td>
<td>&nbsp;</td>
<td>Ranin</td>
<td>L200</td>
<td>&nbsp;</td>
<tr>
<td>Stainless steel mesh, 304, 12x24</td>
<td>&nbsp;</td>
<td>Small Parts</td>
<td>CX-0150-F-01</td>
<td>&nbsp;</td>
<tr>
<td>Glass 5&frac34; inch Pasteur Pipets</td>
<td>&nbsp;</td>
<td>Fisherbrand/Fisher Scientific</td>
<td>13-678-20B</td>
<td>&nbsp;</td>
<tr>
<td>P4 Filter paper</td>
<td>&nbsp;</td>
<td>Fisherbrand/Fisher Scientific</td>
<td>09-803-6F</td>
<td>&nbsp;</td>
<tr>
<td>Rubber bands</td>
<td>&nbsp;</td>
<td>Office Max</td>
<td>A620645</td>
<td>&nbsp;</td>
<tr>
<td>Scotch double-sided tape, &frac12; inch</td>
<td>&nbsp;</td>
<td>Office Max</td>
<td>A8137DM-2</td>
<td>&nbsp;</td>
<tr>
<td>Robert Simmons Expression paint brushes E85 round #2</td>
<td>&nbsp;</td>
<td>Jerry&#x2019;s Artarama</td>
<td>56460</td>
<td>&nbsp;</td>
<tr>
<td>Dumont #5 Forceps High Precision Inox</td>
<td>&nbsp;</td>
<td>Electron Microscopy Services</td>
<td>72701-DZ</td>
<td>&nbsp;</td>
<tr>
<td>Razor blades</td>
<td>&nbsp;</td>
<td>VWR (BD #214010)</td>
<td>55411-050</td>
<td>&nbsp;</td>
<tr>
<td>Halocarbon Oil 27</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>H 8773</td>
<td>&nbsp;</td>
<tr>
<td>Heptane</td>
<td>&nbsp;</td>
<td>Fisher Chemical/Fisher Scientific</td>
<td>H360-1</td>
<td>&nbsp;</td>
<tr>
<td>BD Bacto Agar</td>
<td>&nbsp;</td>
<td>VWR (BD# 214010)</td>
<td>90000-760</td>
<td>&nbsp;</td>
<tr>
<td>Sucrose</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>S7903</td>
<td>&nbsp;</td>
<tr>
<td>p-Hydroxybenzoic acid</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>H5501</td>
<td>&nbsp;</td>
<tr>
<td>Red Star Active Dry Yeast</td>
<td>&nbsp;</td>
<td>LeSaffre</td>
<td>15700</td>
<td>&nbsp;</td>
<tr>
<td>Paper towels, C-fold</td>
<td>&nbsp;</td>
<td>Kleenex</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Heavy duty aluminum foil</td>
<td>&nbsp;</td>
<td>Reynolds Wrap</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Bleach</td>
<td>&nbsp;</td>
<td>Austin&#x2019;s A-1 Commercial</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>100% Apple juice</td>
<td>&nbsp;</td>
<td>Ocean Spray or Tree Top</td>
<td>&nbsp;</td>
<td>&nbsp;</td>		</tbody>
		</table>
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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

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 ...

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 ...

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.