This protocol has been developed through our work within and in collaboration with the laboratories of Paul Martin and Antonio Jacinto. We thank the Bloomington Stock Centre for its excellent service and the Drosophila community for continuing to share fly lines. BS is currently funded by a BBSRC project grant. WW is funded by a Wellcome Trust Career Development Fellowship.

Preparation
<ol>
	<li>Obtain appropriate Drosophila lines containing a hemocyte-specific Gal4 driver (e.g. srp-Gal42) and a genetically encoded fluorescent reporter under UAS control (e.g. uas-GFP). Flies homozygous for srp-Gal4,uas-GMA3 or crq-Gal4,uas-GFP4, 5 are particularly useful for imaging purposes (n.b. GMA is GFP fused to the actin-binding domain of moesin); see below for a discussion of the range of Gal4 drivers and uas constructs available (the Bloomington Stock Centre contains a wide variety).</li>
	<li>Typically genetic crosses are carried out such that mutant alleles are balanced using fluorescent balancers CTG or TTG6, with Gal4 drivers and uas constructs carried on alternative homologous chromosomes. This makes it possible to select homozygous mutants on the basis of absence of CTG/TTG-associated GFP fluorescence (this is done at stage 2.11).</li>
	<li>Amplify stocks and place flies in a laying cage with an apple juice agar plate7. The flies need at least two days to acclimatize to the laying cage before enough embryos begin to be laid. In general twenty flies of each sex should be sufficient to generate enough embryos for imaging but it should be noted that different lines have differing degrees of fertility. We use 55mm Petri dishes that fit into the bottom of a plastic beaker, punctured at its base to allow airflow. The exact means of embryo collection is unimportant, but the timings are critical in order to collect correctly staged embryos.</li>
	<li>Collect embryos from an overnight apple juice agar plate maintained at 25&deg;C or from a timed plate. For the latter we typically allow the flies to lay on a pre-warmed plate for 4 hours, before removing the plate and placing it at 18&deg;C for 15-16 hours prior to mounting of embryos; this provides embryos from late stage 12 through to stage 15 of development. An overnight plate contains a greater diversity of stages but offers the advantage of higher levels of fluorescent reporter expression in hemocytes due to a longer period of time at 25&deg;C as the Gal4-UAS system is temperature sensitive.</li>
</ol>
Procedure
<ol>
<li>Dislodge embryos from the apple juice agar plate using a small amount of water and a soft-tipped paintbrush. Dislodged embryos can be seen easily with the naked eye.</li>
<li>Transfer embryos to a cell strainer (Fisher) or home-made basket7 by pouring water from the apple juice agar plate into the basket held over a beaker to collect waste water.</li>
<li>Repeat step 2.2 until you are satisfied you have enough embryos transferred from the apple juice agar plate.</li>
<li>Wash embryos in cell strainer/basket using water.</li>
<li>Place cell strainer/basket in the Petri dish lid of the apple juice agar plate and add enough neat bleach to suspend embryos in the cell strainer/basket.</li>
<li>Follow dechorionation of the embryos on a dissection microscope under brightfield: dechorionation is complete when the dorsal appendages have dissolved, which should occur within two minutes.</li>
<li>Remove cell strainer/basket containing embryos from the bleach and wash off residual bleach using water. All traces of bleach should be removed before proceeding to step 2.8. One trick to assess whether all the bleach has been removed is to blot off residual water on blue-colored laboratory tissues - if there is residual bleach the blue color will be bleached white/pink.</li>
<li>Blot off remaining water using laboratory tissue/mediwipes applied to the underside of the cell strainer/basket.</li>
<li>Place a droplet of water in a Petri dish lid. With a fine paintbrush, collect all the dechorionated embryos from the embryo basket and resuspend them in the droplet. Next dry the embryos by aspirating water using a micropipette or carefully absorbing it with a laboratory tissue/mediwipes.</li>
<li>Once the embryos have been dried, add a drop of voltalef oil to cover all the embryos. Put a second small drop of oil adjacent to the droplet containing the embryos. N.B. we have been unable to find a UK-based supplier of voltalef oil; halocarbon oil 700 (Sigma) may be used instead. </li>
<li>Under a fluorescent dissection microscope select appropriately staged embryos of the desired genotype using a pair of watchmakers forceps (number 5) from the oil droplet. These forceps should be bent inwards (Figure 1) in order to scoop up the embryos without puncturing their vitelline membrane. Transfer selected embryos to the second oil droplet. It is important that you are able to see fluorescent hemocytes on the dissecting microscope in order to be able to collect good images on the confocal microscope (Figure 2). We typically mount stage 13/14 embryos to image lateral migration of hemocytes on the ventral midline or stage 15 embryos to image the motility of hemocytes following dispersal over the embryo.</li>
<li>Stick two coverslips (18x18mm, thickness 1) to the underside of a Petriperm/Lumox dish (Sarstedt), using 2 small drops of voltalef oil, leaving approximately 1cm between them (Figure 3); these will be used to support a coverslip placed over the embryos, so as not to crush them. Petriperm dishes (50mm diameter) contain a hydrophobic, gas-permeable membrane. We find that the dishes become easier to use once they have been used several times (dishes can be wiped with 70% ethanol and reused).</li>
<li>Under brightfield on the dissection microscope, pick up selected embryos one by one with the bent forceps and line them ventral side up and parallel to the edge of the coverslips (Figure 3). It is possible to align up to 15 embryos in this way, depending on your dexterity and your patience. It is important to manipulate the embryos gently as both the embryos and Petriperm dish membrane are fragile and can be easily ruptured.</li>
<li>Once the embryos are aligned add a small drop of oil and let it spread to form a homogenous layer between the two coverslips. After the oil has spread (this may take a few minutes) check that the embryos are still ventral side up. If the embryos have rolled slightly, reposition them again with the forceps.</li>
<li>Finally, using tweezers (number 3) place a coverslip (18x18mm, thickness 1) over the embryos, resting upon the two previously adhered coverslips. Glue this coverslip to the coverslip supports using nail polish (Figure 3).</li>
<li>Take the Petriperm dish with mounted embryos to the confocal or wide-field microscope and mount the Petriperm dish on the stage using an appropriate adapter. Either an upright or inverted microscope may be used, with the objective lens focusing through the coverslip (as opposed to through the membrane).</li>
</ol>
Representative Result:
This protocol describes how to mount Drosophila embryos for live imaging of hemocytes on the ventral side of the embryo. If done correctly it will be easy to generate either stills or movies of hemocytes. The major determinant is the microscope used to image the hemocytes (in particular the objective lens), but the nature of the images acquired will also depend upon the stage of development, the temperature the embryos were raised at and the Gal4 and uas lines used.
Higher levels of fluorescent protein expression will enable hemocytes to be imaged with greater ease, therefore it is important to be able to see hemocytes when the embryos are at stage 2.11 of the protocol (Figure 2 contains examples of clear hemocytes within embryos, taken with a camera fitted to a dissection microscope). Therefore increased numbers of Gal4 and uas constructs enable a greater signal-to-noise ratio. Furthermore this reduces the need for high laser intensities or increased exposure times when imaging, which in turn enables hemocyte behavior to be followed for longer periods of time.
Very high levels of GFP expression will show up fine details of hemocyte morphology, particularly the thin sheet-like lamellae that surround the circular cell body (Figure 4A-B). Circular regions excluding GFP represent phagosomes (Figure 4A-C). Finger-like filopodia can also be seen emerging from the lamellae (Figure 4B). Two Gal4 drivers remain sufficient to see these processes (Figure 4C), particularly if one or more is srp-Gal4 (see discussion), however slower scanning speeds or greater laser power on the confocal microscope may be required. As expression levels decrease it becomes more difficult to image the protrusions of hemocytes; nonetheless it is still possible to track the migration of hemocytes under these conditions as the cell body remains obvious even when protrusions are less clear (Figure 4D).
At earlier stages of development (up to stage 13) hemocytes migrate in close contact to one another and it is often hard to distinguish individual cells. By the end of stage 13 hemocytes have formed a single line down the ventral midline (Figure 5A), then, becoming more motile, migrate laterally to the edges of the ventral nerve cord (Figure 5B). The actin cytoskeleton within the dynamic protrusions of hemocytes can be directly observed through expression of GMA (Figure 5C) or cherry-moesin.
Mounting the embryos in this way allows gas exchange and prevents dehydration and embryos remain viable following imaging. If the embryo is damaged during mounting it is generally obvious as the embryo content will leak through its vitelline membrane. If an embryo does start to dehydrate then this can often been seen by deformations in the vitelline membrane. Occasionally an embryo will roll during the course of a timelapse movie, however this only tends to be problematic for longer timescale movies. Lastly, mounting several embryos at once gives the experimentalist the best chance of obtaining an embryo in the perfect orientation for their experiment.
<img src="/files/ftp_upload/1696/1696fig1.jpg" alt="Figure 1" /> Figure 1. Forceps for manipulation of dechorionated embryos. The tips of watchmakers forceps (size number 5) should be bent inwards in order to fashion a tool to scoop up embryos as shown here. The outer surface of the bent region is also useful to manipulate embryos when positioning on the Petriperm membrane as they possess no sharp edges that could puncture the embryo.
<img src="/files/ftp_upload/1696/1696fig2.jpg" alt="Figure 2" /> Figure 2. Representative images of embryos that will yield good live imaging results. Images of dechorionated embryos in voltalef oil (at stage 2.11 of the protocol) taken on a fluorescent dissecting microscope. Lateral views of stage 13 (A) and stage 15 (B) srp-Gal4,uas-GFP;crq-Gal4,uas-GFP embryos. Lateral view of a stage 15 srp-Gal4,uas-GFP/+;crq-Gal4,uas-GFP/uas-N17Rac embryo (C) in which hemocytes have failed to migrate out of the head, demonstrating what embryos look like when hemocytes are not apparent along their migratory routes. Lateral view of a stage 17 srp-Gal4,uas-GFP;crq-Gal4,uas-GFP embryo showing the convoluted structure of the gut at this stage of development (D); the onset of muscle contraction prevents live imaging of embryos beyond this stage of development. Ventral views of stage 13 (E) and stage 14 (F) srp-Gal4,uas-red stinger embryos showing dispersal of hemocytes with fluorescently labeled nuclei. Observation of hemocytes by fluorescence at stage 2.11 of the protocol is a prerequisite to obtain excellent images; anterior is to the right for all images.
<img src="/files/ftp_upload/1696/1696fig3.jpg" alt="Figure 3" /> Figure 3. Mounting of embryos on a Petriperm/Lumox dish. Two 18x18mm coverslips (thickness 1) are stuck to the bottom face of the Petriperm dish using a small drop of oil, separated by about 1cm as shown. Embryos are then lined up ventral side up with their long (anterior-posterior) axis parallel to the edges of the coverslips and covered with a small drop of oil. Once the oil has spread to fill the gap between the two coverslips a third coverslip (18x18mm thickness 1) is gently placed on top of the oil-covered embryos using the two previously adhered coverslips as a bridge to prevent the embryos from being squashed. This coverslip is then glued to the two coverslip bridges using two small drops of nail polish. Once set, the embryos can be imaged on an upright or inverted microscope with the objective lens focusing down through the coverslip (as opposed to through the Petriperm membrane).
<img src="/files/ftp_upload/1696/1696fig4.jpg" alt="Figure 4" /> Figure 4. Representative results from live imaging of GFP labeled hemocytes. Z-projections of hemocytes on the ventral side of a stage 14 srp-Gal4,uas-GFP;crq-Gal4,uas-GFP embryo (A-B). (A) is a lower magnification image such as used to monitor hemocyte developmental migrations in timelapse movies. (B) is a higher magnification still of hemocytes on the ventral midline, showing fine details of their morphology. (C) is a single 1 m slice of hemocytes on the ventral midline in a stage 14 srp-Gal4,uas-GFP/+;crq-Gal4,uas-GFP/+ embryo, revealing that lower copy numbers of Gal4 drivers and uas constructs are also sufficient to generate good images. (D) displays a z-projection of hemocytes in a stage 14 crq-Gal4,uas-GFP embryo. Here hemocyte protrusions are less obvious due to lower expression of GFP but it is still possible to make movies and track hemocyte migration with this combination of Gal4 driver and uas construct. Images were taken on a Leica LSM510 confocal microscope; anterior is up in all images; the rings at the periphery of images are caused by vitelline membrane autofluorescence.
<img src="/files/ftp_upload/1696/1696fig5.jpg" alt="Figure 5" /> Figure 5. Representative results from live imaging of GMA expressing hemocytes. Z-projections of hemocytes on the ventral midline of stage 13 (A) and stage 14 (B) srp-Gal4,uas-GMA embryos, taken from timelapse movies to show developmental migrations of hemocytes. Detailed information on actin dynamics can be obtained by higher magnification imaging of GMA expressing hemocytes (C). GMA consists of GFP fused to the actin-binding domain of moesin and labels actin filaments. Anterior is up in all images; images were taken on a confocal microscope.

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P. <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+morphogenetic+movements+in+Drosophila+using+Gal4-UAS-driven+expression+of+GFP+fused+to+the+actin-binding+domain+of+moesin.">Real-time imaging of morphogenetic movements in Drosophila using Gal4-UAS-driven expression of GFP fused to the actin-binding domain of moesin.</a> Genesis. 34, 146-151 (2002).</li><li>Stramer, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live+imaging+of+wound+inflammation+in+Drosophila+embryos+reveals+key+roles+for+small+GTPases+during+in+vivo+cell+migration.">Live imaging of wound inflammation in Drosophila embryos reveals key roles for small GTPases during in vivo cell migration.</a> J Cell Biol. 168, 567-573 (2005).</li><li>Wood, W., Jacinto, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+melanogaster+embryonic+haemocytes:+masters+of+multitasking.">Drosophila melanogaster embryonic haemocytes: masters of multitasking.</a> Nat Rev Mol Cell Biol. 8, 542-551 (2007).</li><li>Halfon, M. 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The most important elements of this procedure are the selection of healthy embryos with clearly labeled hemocytes and to mount them carefully without damaging them. Once the embryos are in the halocarbon oil they are resistant to dehydration and once mounted can be imaged for several hours. In our hands we can image hemocytes for three hours, with negligible dehydration of the embryo or obvious photo-damage, taking a z-stack of images every three minutes on our Zeiss LSM510 confocal microscope with a 40X objective. Since...

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		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
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<td>Cell strainer</td>
<td>&nbsp;</td>
<td>BD Falcon</td>
<td>352350</td>
<td>70&mu;m pores</td>
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<td>Halcarbon oil 700</td>
<td>&nbsp;</td>
<td>Sigma</td>
<td>H8898</td>
<td>&nbsp;</td>
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<td>Lumox/Petriperm dish</td>
<td>&nbsp;</td>
<td>Sarstedt</td>
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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.

Watch this Scientific Journal Video about Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations at JoVE.com

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.

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 ...

live-imaging-of-drosophila-melanogaster-embryonic-hemocyte-migrations

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JoVE Journal

Biology

15.6K visualizações.  University of Bath. Hemócitos Drosophila dispersam sobre a totalidade do embrião em desenvolvimento. Este protocolo demonstra como montar e imagem essas migrações usando embriões com hemócitos fluorescente etiquetado.

Vídeo: Live Imaging Of Drosophila melanogaster Embryonic Hemocyte Migrations

Dissection of Larval CNS in Drosophila Melanogaster

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to examine the expression of various novel and marker proteins.  Staining of whole larvae is impractical because the tough cuticle prevents antibodies from penetrating inside the body cavity.  In order to stain these tissues it is necessary to dissect the animal prior to fixing and staining.  In this article we demonstrate how to dissect Drosophila larvae without damaging the CNS.  Begin by tearing the larva in half with a pair of fine forceps, and then turn the cuticle "inside-out" to expose the CNS.  If the dissection is performed carefully the CNS will remain attached to the cuticle.  We usually keep the CNS attached to the cuticle throughout the fixation and staining steps, and only completely remove the CNS from the cuticle just prior to mounting the samples on glass slides.  We also show some representative images of a larval CNS stained with Eve, a transcription factor expressed in a subset of neurons in the CNS.  The article concludes with a discussion of some of the practical uses of this technique and the potential difficulties that may arise.

In this article we demonstrate how to dissect the central nervous system from third instar Drosophila larvae.

The central nervous system (CNS) of Drosophila larvae is complex and poorly understood.  One way to investigate the CNS is to use immunohistochemistry to ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes in cell surface area and cell height, and where cells undergo mitosis and migrate. Furthermore, epithelial cells can also regulate morphogenetic movements in adjacent tissues1. A traditional method to study epithelial cells and tissues involve chemical fixation and histological methods to determine cell morphology or localization of particular proteins of interest. These approaches continue to be useful and provide "snapshots" of cell shapes and tissue architecture, however, much remains to be understood about how cells acquire specific shapes, how various proteins move or localize to specific positions, and what paths cells follow toward their final differentiated fate. High resolution live imaging complements traditional methods and also allows more direct investigation into the dynamic cellular processes involved in the formation, maintenance, and morphogenesis of multicellular epithelial sheets. Here we demonstrate experimental methods from the isolation of animal cap tissues from Xenopus laevis embryos to confocal imaging of epithelial cells and simple measurement approaches that together can augment molecular and cellular studies of epithelial morphogenesis.

Live-cell Imaging and Quantitative Analysis of Embryonic Epithelial Cells in Xenopus laevis

Xenopus embryonic epithelia are an ideal model system to study cell behaviors such as polarity development and shape change during epithelial morphogenesis. Traditional histology of fixed samples is increasingly being complemented by live-cell confocal imaging. Here we demonstrate methods to isolate frog tissues and visualize live epithelial cells and their cytoskeleton using live-cell confocal microscopy.

Embryonic epithelial cells serve as an ideal model to study morphogenesis where multi-cellular tissues undergo changes in their geometry, such as changes ...

Dissection of Oenocytes from Adult Drosophila melanogaster

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides protection against desiccation and other environmental challenges. Several of these cuticular hydrocarbon (CHC) compounds also function as semiochemical signals, and as such mediate pheromonal communications between members of the same species, or in some instances between different species, and influence behavior. Specialized cells referred to as oenocytes are regarded as the primary site for CHC synthesis. However, relatively little is known regarding the involvement of the oenocytes in the regulation of the biosynthetic, transport, and deposition pathways contributing to CHC output. Given the significant role that CHCs play in several aspects of insect biology, including chemical communication, desiccation resistance, and immunity, it is important to gain a greater understanding of the molecular and genetic regulation of CHC production within these specialized cells. The adult oenocytes of D. melanogaster are located within the abdominal integument, and are metamerically arrayed in ribbon-like clusters radiating along the inner cuticular surface of each abdominal segment. In this video article we demonstrate a dissection technique used for the preparation of oenocytes from adult D. melanogaster. Specifically, we provide a detailed step-by-step demonstration of (1) how to fillet prepare an adult Drosophila abdomen, (2) how to identify the oenocytes and discern them from other tissues, and (3) how to remove intact oenocyte clusters from the abdominal integument. A brief experimental illustration of how this preparation can be used to examine the expression of genes involved in hydrocarbon synthesis is included. The dissected preparation demonstrated herein will allow for the detailed molecular and genetic analysis of oenocyte function in the adult fruit fly.

Dissection of Oenocytes from Adult Drosophila melanogaster

In insects, the oenocytes produce cuticular hydrocarbon compounds. These compounds protect against desiccation and facilitate chemical communication. Here we demonstrate a dissection technique used to isolate the oenocytes from adult Drosophila melanogaster, and illustrate how this preparation can be utilized to study genes involved in hydrocarbon synthesis.

In Drosophila melanogaster, as in other insects, a waxy layer on the outer surface of the cuticle, composed primarily of hydrocarbon compounds, provides ...

Isolation of Drosophila melanogaster Testes

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline interactions. Testes are typically isolated from adult males 0-3 days after eclosion from the pupal case. The testes of wild-type flies are easily distinguished from other tissues because they are yellow, but the testes of white mutant flies, a common genetic background for laboratory experiments are similar in both shape and color to the fly gut. Performing dissection on a glass microscope slide with a black background makes identifying the testes considerably easier. Testes are removed from the flies using dissecting needles. Compared to protocols that use forceps for testes dissection, our method is far quicker, allowing a well-practiced individual to dissect testes from 200-300 wild-type flies per hour, yielding 400-600 testes. Testes from white flies or from mutants that reduce testes size are harder to dissect and typically yield 200-400 testes per hour.

Isolation of Drosophila melanogaster Testes

Drosophila melanogaster testes can be rapidly and efficiently isolated from adult males using dissecting needles. With practice, one can readily isolate in one or two days an amount of testes sufficient for the analysis of DNA or RNA by high throughput sequencing or more traditional molecular biology methods or of protein for antibody- or enzyme-based assays.

The testes of Drosophila melanogaster provide an important model for the study of stem cell maintenance and differentiation, meiosis, and soma-germline ...

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, ...

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.

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.

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

Measurement of Lifespan in Drosophila melanogaster

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced physical performance and increased risk of disease. Individual aging is manifest at the population level as an increase in age-dependent mortality, which is often measured in the laboratory by observing lifespan in large cohorts of age-matched individuals. Experiments that seek to quantify the extent to which genetic or environmental manipulations impact lifespan in simple model organisms have been remarkably successful for understanding the aspects of aging that are conserved across taxa and for inspiring new strategies for extending lifespan and preventing age-associated disease in mammals.
The vinegar fly, Drosophila melanogaster, is an attractive model organism for studying the mechanisms of aging due to its relatively short lifespan, convenient husbandry, and facile genetics. However, demographic measures of aging, including age-specific survival and mortality, are extraordinarily susceptible to even minor variations in experimental design and environment, and the maintenance of strict laboratory practices for the duration of aging experiments is required. These considerations, together with the need to practice careful control of genetic background, are essential for generating robust measurements. Indeed, there are many notable controversies surrounding inference from longevity experiments in yeast, worms, flies and mice that have been traced to environmental or genetic artifacts1-4. In this protocol, we describe a set of procedures that have been optimized over many years of measuring longevity in Drosophila using laboratory vials. We also describe the use of the dLife software, which was developed by our laboratory and is available for download (<a href="http://sitemaker.umich.edu/pletcherlab/software">http://sitemaker.umich.edu/pletcherlab/software</a>). dLife accelerates throughput and promotes good practices by incorporating optimal experimental design, simplifying fly handling and data collection, and standardizing data analysis. We will also discuss the many potential pitfalls in the design, collection, and interpretation of lifespan data, and we provide steps to avoid these dangers.

Measurement of Lifespan in Drosophila melanogaster

Drosophila melanogaster is a powerful model organism for exploring the molecular basis of longevity regulation. This protocol will discuss the steps involved in generating a reproducible, population-based measurement of longevity as well as potential pitfalls and how to avoid them.

Aging is a phenomenon that results in steady physiological deterioration in nearly all organisms in which it has been examined, leading to reduced ...

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  ...

Imaging Through the Pupal Case of Drosophila melanogaster

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled analysis of cell and developmental biology from a variety of methodological angles. Live imaging is an emerging method for observing dynamic cell processes, such as cell division or cell motility. Having isolated mutations in uncharacterized putative cell cycle proteins it became essential to observe mitosis in situ using live imaging. Most live imaging studies in Drosophila have focused on the embryonic stages that are accessible to manipulation and observation because of their small size and optical clarity. However, in these stages the cell cycle is unusual in that it lacks one or both of the gap phases. By contrast, cells of the pupal wing of Drosophila have a typical cell cycle and undergo a period of rapid mitosis spanning about 20 hr of pupal development. It is easy to identify and isolate pupae of the appropriate stage to catch mitosis in situ. Mounting intact pupae provided the best combination of tractability and durability during imaging, allowing experiments to run for several hours with minimal impact on cell and animal viability. The method allows observation of features as small as, or smaller than, fly chromosomes. Adjustment of microscope settings and the details of mounting, allowed extension of the preparation to visualize membrane dynamics of adjacent cells and fluorescently labeled proteins such as tubulin. This method works for all tested fluorescent proteins and can capture submicron scale features over a variety of time scales. While limited to the outer 20 &#181;m of the pupa with a conventional confocal microscope, this approach to observing protein and cellular dynamics in pupal tissues in vivo may be generally useful in the study of cell and developmental biology in these tissues.

Imaging Through the Pupal Case of Drosophila melanogaster

This paper demonstrates the use of a fast scanning confocal microscope to image cell behavior directly through the puparium. By leaving the pupal case intact, this method allows observation and measurement of dynamic cell processes at a stage of Drosophila development that is difficult to study directly.

The longstanding use of Drosophila as a model for cell and developmental biology has yielded an array of tools. Together, these techniques have enabled ...