This work was supported by grant GM096076 from the NIH AREA program.

1. Preparing Flies for Dissection
<ol>
 <li>Anesthetize healthy flies of the desired genotype (for example, expressing a GFP-labeled protein) that are less than one week old. Select 10-15 large females with rounded cream-colored abdomens.</li>
 <li>Transfer the selected flies and a few (3-5) males to a vial containing new lightly yeasted food and allow the flies to fatten for two days at 25 &deg;C. Refer to Weil et al.6 for details of Drosophila preparation. Fattening the flies ensures that a variety of stages, especially stages 8-12, will be available for imaging. Poorly fattened or unfattened flies usually contain only very early stages (1-6) and mature (stage 14) oocytes.</li>
</ol>
2. Preparation of Oocytes for Imaging Using an Upright Compound Microscope
<ol>
 <li>Prepare a standard glass slide by affixing two coverslips to the slide, approximately 1 cm apart, using silicone grease. Only use enough grease to ensure a good seal between the coverslip and slide. Pressing down firmly on each the coverslip will spread the grease thinly and evenly. Add some halocarbon oil 27 (Sigma) to the juncture between the coverslips and slide. They will serve as spacers to prevent injuring isolated oocytes in subsequent steps. No. 1.5 coverslips work well, with an average thickness (0.16-0.18 mm) that matches that of late stage oocytes.</li>
 <li>Place 2-3 drops of halocarbon oil 27 onto the surface of a 22 mm2 coverslip. To obtain the healthiest oocytes possible, an individual female is removed from the food vial using mouth pipetting and gently deposited into the halocarbon oil without first anesthetizing.</li>
 <li>Using sharp dissecting forceps (such as Dumont #5), pin the fly against the coverslip and pull off the tip of the abdomen. Remove the ovaries by dragging them along the surface of the coverslip, under the oil. This will promote adhesion of the oocytes to the coverslip.</li>
 <li>Gently tease apart the ovaries, looking for oocytes that are at least stage 10B of oogenesis. Oocytes at this stage can be identified by looking for egg chambers in which the oocyte occupies at least 50-60% of the total volume of the egg chamber. At this point in oogenesis, the follicle cells overlying the oocyte have become columnar, and surround the oocyte on all sides except for a small region where connections to the nurse cells remain. Use the forceps to remove individual oocytes from the ovariole sheath.</li>
 <li>Proceed, using 1-3 females, until 5-8 undamaged oocytes have been isolated on the coverslip. Remove any unuseable tissue.</li>
 <li>Carefully invert the coverslip containing the oocytes onto the pre-prepared slide so that the oocytes are positioned between the two spacers. Do not press down on the inverted coverslip as that will damage the oocytes. If necessary, add a small amount of halocarbon oil to the edges of the "sandwich" to ensure the space is filled. (Optional: Let slide rest for a several minutes to allow for settling of coverslip.) The slide is now ready for imaging.</li>
</ol>
3. Preparation of Oocytes for Imaging Using an Inverted Compound Microscope
<ol>
 <li>Oocytes are dissected essentially as described in section 2, except that small Petri dishes with a glass bottom insert (MatTek, 35 mm) are used instead of coverslips. It is unnecessary to cover the specimen, and after dissection, tissue can be imaged directly in the dishes, but care must be taken to ensure the oocytes remain covered with oil and are not allowed to dry out.</li>
</ol>
4. Live Imaging
<ol>
 <li>Bring an oocyte into focus using DIC or phase contrast optics and examine the oocyte for any signs of damage, such as leaking of cytoplasm, or bulging of follicle cells. Image only oocytes that appear healthy and undamaged.</li>
 <li>Streaming can be monitored directly without the need for GFP labeling, since yolk vesicles in the cytoplasm are autofluorescent7. This signal can be captured in the FITC range of the scope but will likely require higher gain settings than those used for GFP-labeled proteins.</li>
 <li>The best images of streaming are obtained from optical sections that are just below the surface of the oocyte that is resting against the glass coverslip/Petri glass. Here the oocyte is slightly flattened, making for a better plane of focus. Images can be captured at 200X - 1,000X. Using the arrangement described here, air objectives should be used, since the coverslip forms a barrier between the oocyte and objective. This is optimal because it greatly minimizes drift and movement of the sample.</li>
 <li>Once a region of interest is in focus, turn off the brightfield optics and switch to confocal imaging. Using the fast scan/preview function of the software, adjust focus and gain as necessary.</li>
 <li>Settings for capture will vary from microscope to microscope, but general parameters are as follows: Use an excitation wavelength of ~488 nm and an emission wavelength of ~ 515 nm. Set up Z-axis capturing so that the Z-axis remains constant, with a lag of 10 sec between frames. Capture a test sequence of 5-10 frames to ensure the oocyte is stationary and the plane of focus is appropriate.</li>
 <li>A typical time lapse sequence will capture between 20 - 50 frames, and can be reviewed immediately after completion to assess video quality. The process can be repeated for other cells in the slide, or additional females can be dissected as needed.</li>
</ol>
5. Troubleshooting and Common Problems
<ol>
 <li>Drift - When using an upright microscope, cells can sometimes drift due to spreading of oil. This can be minimized by using only enough oil to cover the area under the coverslip. If problems persist, use less silicone grease to affix spacers to the slide.</li>
</ol>
Note: Some confocal software packages allow for cell tracking, but in general it is better to abandon imaging of drifting oocytes and instead use another specimen.
<ol start="2">
 <li>No signs of streaming - If streaming is not evident, it is likely due to one of two causes; either the focal plane for image capture is not correct (usually too deep into the interior of the oocyte) or the oocyte is damaged. Time and practice can minimize both errors.</li>
</ol>

<ol>
	<li>Becalska, A. N., Gavis, E. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lighting+up+mRNA+localization+in+Drosophila+oogenesis.">Lighting up mRNA localization in Drosophila oogenesis.</a> Dev. 136, 2493-2503 (2009).</li><li>He, L., Wang, X., Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shining+light+on+Drosophila+oogenesis:+Live+imaging+of+egg+development.">Shining light on Drosophila oogenesis: Live imaging of egg development.</a> Curr. Opin. Gen. Dev. 21, 612-619 (2011).</li><li>King, R. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Ovarian development Drosophila melanogaster. , (1970).</li><li>Spradling, A. C., Bate, M., Martinez-Arias, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Developmental+genetics+of+oogenesis.">Developmental genetics of oogenesis.</a> The Development of Drosophila melanogaster. 1, 1-70 (1993).</li><li>Serbus, L. R., Cha, B. -. J., Theurkauf, W. E., Saxton, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynein+and+the+actin+cytoskeleton+control+kinesin-driven+cytoplasmic+streaming+in+Drosophila+oocytes.">Dynein and the actin cytoskeleton control kinesin-driven cytoplasmic streaming in Drosophila oocytes.</a> Dev. 132, 3743-3752 (2005).</li><li>Weil, T. T., Parton, R. M., Davis, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparing+Individual+Drosophila+Egg+Chambers+for+Live+Imaging.">Preparing Individual Drosophila Egg Chambers for Live Imaging.</a> J. Vis. Exp. (60), e3679 (2012).</li><li>Theurkauf, W. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Premature+microtubule-dependent+cytoplasmic+streaming+in+cappuccino+and+spire+mutant+oocytes.">Premature microtubule-dependent cytoplasmic streaming in cappuccino and spire mutant oocytes.</a> Science. 265, 2093-2096 (1998).</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> PNAS. 98, 15050-15055 (2001).</li><li>R&#246;per, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rtnl1+is+enriched+in+a+specialized+germline+ER+that+associates+with+ribonucleoprotein+granule+components.">Rtnl1 is enriched in a specialized germline ER that associates with ribonucleoprotein granule components.</a> J. Cell Sci. 120, 1081-1092 (2007).</li><li>Pokrywka, N. J., Payne-Tobin, A., Raley-Susman, K. M., Swartzman, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microtubules,+the+ER+and+Exu:+New+associations+revealed+by+analysis+of+mini+spindles+mutations.">Microtubules, the ER and Exu: New associations revealed by analysis of mini spindles mutations.</a> Mech. Dev. 126, 289-300 (2009).</li><li>Petri, W. H., Mindrinos, M. N., Lombard, M. F., Margaritis, L. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+vitro+development+of+the+Drosophila+chorion.ih+a+chemically+defined+organ+culture+medium.">In vitro development of the Drosophila chorion.ih a chemically defined organ culture medium.</a> Wilhelm Roux's Arch., Devl. Biol. 186, 351-362 (1979).</li></ol>

I have nothing to disclose.

Drosophila oocytes are a versatile model system for addressing a variety of questions related to the function and organization of cells and subcellular components. A complete understanding of protein function requires the monitoring of both spatial and temporal aspects of protein behavior. Advances in both imaging systems and the genetic tools available to Drosophilists make it possible for even small labs to perform high quality imaging experiments in living oocytes. Here I outline a protocol to analyze the behavior of GFP-tagged or autofluorescent proteins and structures during mid- to late-oogenesis that can be used in conjunction with a variety of genetic backgrounds to probe protein function.
In this protocol I describe the process by which oocytes are prepared for imaging and the basic confocal settings that will enable acquisition of time lapse video of fluorescent proteins and structures. Additional details on oocyte preparation are described by Weil et al.6 A wide variety of fly strains expressing proteins that have been endogenously tagged via exon-trapping are available8, and infinitely more protein variants can be engineered and reintroduced to flies.
In working with living tissue, the health of the specimens is of paramount importance. Egg chambers from stage 10B can complete oogenesis in an essentially autonomous fashion11, making them ideal for short term imaging studies. Care must be taken at several key steps to get high quality data. During dissection it is important to manipulate ovaries and oocytes as little as possible, and to minimize the amount of time between dissection and the completion of imaging. Ideally, a small dissection station should be located in the same room that houses the confocal scope, and only a few egg chambers at a time should be imaged. The number recommended here (5-8) ensures that dissection time is minimized while maximizing the likelihood of obtaining good quality images. I recommend capturing a short series of images to assess image quality and confirm that capture settings are optimized, before capturing a longer time lapse sequence. Most software allows individual frames to be saved, and these images can then be analyzed in a variety of ways using ImageJ12 or other software.

<table border="1">
 <tbody>
 <tr>
 <td>Name</td>
 <td>Company</td>
 <td>Catalog number</td>
 <td>Comments</td>
 </tr>
 <tr>
 <td>Halocarbon oil 27</td>
 <td>Sigma-Aldrich</td>
 <td>H8773-100ML </td>
 <td></td>
 </tr>
 <tr>
 <td>#5SF Forceps </td>
 <td>Fine Science Tools</td>
 <td>11252-00 </td>
 <td></td>
 </tr>
 <tr>
 <td>Silicon grease</td>
 <td>Sigma Aldrich</td>
 <td>Z273554-1EA </td>
 <td></td>
 </tr>
 <tr>
 <td>Glass-bottom Petri dishes</td>
 <td>MatTek</td>
 <td>P35G-0-20-C </td>
 <td></td>
 </tr>
 </tbody>
</table>

live imaging of gfp-labeled proteins in drosophila oocytes

The Drosophila oocyte has been established as a versatile system for investigating fundamental questions such as cytoskeletal function, cell organization, and organelle structure and function. The availability of various GFP-tagged proteins means that many cellular processes can be monitored in living cells over the course of minutes or hours, and using this technique, processes such as RNP transport, epithelial morphogenesis, and tissue remodeling have been described in great detail in Drosophila oocytes1,2.
The ability to perform video imaging combined with a rich repertoire of mutants allows an enormous variety of genes and processes to be examined in incredible detail. One such example is the process of ooplasmic streaming, which initiates at mid-oogenesis3,4. This vigorous movement of cytoplasmic vesicles is microtubule and kinesin-dependent5 and provides a useful system for investigating cytoskeleton function at these stages.
Here I present a protocol for time lapse imaging of living oocytes using virtually any confocal microscopy setup.

Imaging of GFP-labeled proteins will vary depending on how strongly the tagged protein is expressed. A mid-stage oocyte expressing reticulon-like 1 (Rtnl-1), exon tagged with GFP8,9 produces strong signal. Rtnl-1 appears to co-localize with the long microtubules present during ooplasmic streaming10 (Figure 1A). Rtnl-1 is a good marker for observing streaming, and is also an indicator of microtubule organization in late stage oocytes. 
Ooplasmic streaming in a wildtype egg chamber is evident as noticeable movement of autofluorescent yolk vesicles when using a capture rate of one frame every 10 sec (Figure 1B). A maximum projection of five frames can be generated to mark the path of moving vesicles.
<img alt="Figure 1" src="/files/ftp_upload/50044/50044fig1.jpg" /> 
 Figure 1. Live imaging of both autofluorescent and GFP-tagged structures. A) A single time-lapse image of a stage 11 oocyte expressing Rtnl1-GFP at 400X. Rtnl1-GFP fibers move with a wave-like motion in streaming oocytes. B) A five frame maximum projection of a stage 10B egg chamber in which the autofluorescent yolk vesicles were imaged every 10 sec for 50 sec total, at 400X magnification. Scale bar = 30 &mu;m.

Watch this Scientific Journal Video about Live Imaging of GFP-labeled Proteins in Drosophila Oocytes at JoVE.com

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

The  Drosophila oocyte has been established as a versatile system for investigating  fundamental questions such as cytoskeletal function, cell ...

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11.2K Views.  Vassar College. A protocol for live imaging of GFP-tagged proteins or autofluorescent structures in individual Drosophila oocytes is described.

Video: Live Imaging of GFP-labeled Proteins in Drosophila Oocytes

Live Imaging of Glial Cell Migration in the Drosophila Eye Imaginal Disc

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons. While recent progress has been made to describe the live migration of glial cells in the developing pupal wing (1), studies of Drosophila glial cell migration have typically involved the examination of fixed tissue. Live microscopic analysis of motile cells offers the ability to examine cellular behavior throughout the migratory process, including determining the rate of and changes in direction of growth. Paired with use of genetic tools, live imaging can be used to determine more precise roles for specific genes in the process of development. Previous work by Silies et al. (2) has described the migration of glia originating from the optic stalk, a structure that connects the developing eye and brain, into the eye imaginal disc in fixed tissue. Here we outline a protocol for examining the live migration of glial cells into the Drosophila eye imaginal disc. We take advantage of a Drosophila line that expresses GFP in developing glia to follow glial cell progression in wild type and in mutant animals.

Here we describe a protocol to examine the migration of glial cells into the developing Drosophila eye using live microscopic analysis paired with GFP tagged glial cells.

Glial cells of both vertebrate and invertebrate organisms must migrate to final target regions in order to ensheath and support associated neurons.  While ...

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 ...

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 ...

Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest ...

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three preparations of the brain and visual system that cover the range from the developing eye disc-brain complex in the developing pupae to individual eye and brain dissection from adult flies. All protocols are optimized for the live culture of the preparations. However, we also present the conditions for fixed tissue immunohistochemistry where applicable. Finally, we show live imaging conditions for these preparations using conventional and resonant 4D confocal live imaging in a perfusion chamber. Together, these protocols provide a basis for live imaging on different time scales ranging from functional intracellular assays on the scale of minutes to developmental or degenerative processes on the scale of many hours.

Preparation of Developing and Adult Drosophila Brains and Retinae for Live Imaging

This protocol describes three Drosophila preparations: 1) adult brain dissection, 2) adult retina dissection and 3) developing eye disc- brain complexes dissection. Emphasis is laid on special preparation techniques and conditions for live imaging, although all preparations can be used for fixed tissue immunohistochemistry.

The Drosophila brain and visual system are widely utilized model systems to study neuronal development, function and degeneration. Here we show three ...

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.

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.

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

Preparing Individual Drosophila Egg Chambers for Live Imaging

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, cell differentiation and organogenesis 1. Correct preparation of the experimental samples is a crucial, often neglected, step. The goal of preparation is to ensure physiological relevance and to establish optimal imaging conditions. To maintain tissue viability, it is critical to avoid dehydration, hypoxia, overheating or medium deterioration 2. 
 The Drosophila egg chamber is a well established system for examining questions relating, but not limited, to body patterning, mRNA localization and cytoskeletal organization 3,4. For early- and mid-stage egg chambers, mounting in halocarbon oil is good for survival in that it allows free diffusion of oxygen, prevents dehydration and hypoxia and has superb optical properties for microscopy. Imaging of fluorescent proteins is possible through the introduction of transgenes into the egg chamber or physical injection of labeled RNA, protein or antibodies 5-7. For example, addition of MS2 constructs to the genome of animals enables real time observation of mRNAs in the oocyte 8. These constructs allow for in vivo labeling of mRNA through utilization of the MS2 bacteriophage RNA stem loop interaction with its coat protein 9. 
 Here, we present a protocol for the extraction of ovaries as well as isolating individual ovarioles and egg chambers from the female Drosophila. For a detailed description of Drosophila oogenesis see Allan C. Spradling (1993, reprinted 2009) 10.

Preparing Individual Drosophila Egg Chambers for Live Imaging

The Drosophila egg chamber is an excellent model for studying the mechanisms of mRNA localization. In order to capture the dynamic events that underpin the processes of localization, rapid high resolution imaging of live tissue is required. Here, we present a protocol for dissection and imaging of live samples with minimal disruption.

Live cell imaging is an important technique applied to a number of Drosophila tissues used as models to investigate topics such as axis specification, ...

Two- and Three-Dimensional Live Cell Imaging of DNA Damage Response Proteins

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate mutations and chromosomal aberrations1. To prevent this trauma from catalyzing genomic instability, it is crucial for cells to detect DSBs, activate the DNA damage response (DDR), and repair the DNA. When stimulated, the DDR works to preserve genomic integrity by triggering cell cycle arrest to allow for repair to take place or force the cell to undergo apoptosis. The predominant mechanisms of DSB repair occur through nonhomologous end-joining (NHEJ) and homologous recombination repair (HRR) (reviewed in2). There are many proteins whose activities must be precisely orchestrated for the DDR to function properly. Herein, we describe a method for 2- and 3-dimensional (D) visualization of one of these proteins, 53BP1.
The p53-binding protein 1 (53BP1) localizes to areas of DSBs by binding to modified histones3,4, forming foci within 5-15 minutes5. The histone modifications and recruitment of 53BP1 and other DDR proteins to DSB sites are believed to facilitate the structural rearrangement of chromatin around areas of damage and contribute to DNA repair6. Beyond direct participation in repair, additional roles have been described for 53BP1 in the DDR, such as regulating an intra-S checkpoint, a G2/M checkpoint, and activating downstream DDR proteins7-9. Recently, it was discovered that 53BP1 does not form foci in response to DNA damage induced during mitosis, instead waiting for cells to enter G1 before localizing to the vicinity of DSBs6. DDR proteins such as 53BP1 have been found to associate with mitotic structures (such as kinetochores) during the progression through mitosis10.
In this protocol we describe the use of 2- and 3-D live cell imaging to visualize the formation of 53BP1 foci in response to the DNA damaging agent camptothecin (CPT), as well as 53BP1's behavior during mitosis. Camptothecin is a topoisomerase I inhibitor that primarily causes DSBs during DNA replication. To accomplish this, we used a previously described 53BP1-mCherry fluorescent fusion protein construct consisting of a 53BP1 protein domain able to bind DSBs11. In addition, we used a histone H2B-GFP fluorescent fusion protein construct able to monitor chromatin dynamics throughout the cell cycle but in particular during mitosis12. Live cell imaging in multiple dimensions is an excellent tool to deepen our understanding of the function of DDR proteins in eukaryotic cells.

This protocol describes a method for visualizing a DNA double-strand break signaling protein activated in response to DNA damage as well as its localization during mitosis.

Double-strand breaks (DSBs) are the most deleterious DNA lesions a cell can encounter. If left unrepaired, DSBs harbor great potential to generate ...

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  ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...