The authors would like to thank Michael Anderson for establishing in the Lee lab these accepted methods for studying spermatogenesis with expert advice from Karen Hales. H. Oda and Y. Akiyama-Oda generously provided the &#947;-tubulin-GFP fly stock. This work was supported by an NIH R01 grant to L.A.L. (GM074044).

1.&#160;Testes Dissection
<ol>
	<li>Anesthetize flies in a bottle or vial using a stream of CO2 and transfer to a fly pad.</li>
	<li>Sort flies under a dissecting microscope using a small paintbrush, and collect an appropriate number (depending on the experiment) of Drosophila males of the desired genotypes. Young males (0-2 days old) are ideal for examining cells throughout the earlier stages of spermatogenesis (e.g. spermatogonia, spermatocytes, and early post-meiotic spermatids), whereas slightly older males (2-5 days old) are ideal for examining cells in the final stages of spermatogenesis (in particular, mature sperm).</li>
	<li>Use forceps to remove the wings from each fly (to prevent the flies from floating in liquid during dissection).</li>
	<li>Add ~500 ml of phosphate-buffered saline (130 mM NaCl, 7 mM Na2HPO4, 3 mM NaH2PO4; PBS) in a drop to a silicone-coated dissection dish on a black background. Other aqueous solutions have been successfully used for testes dissection3.</li>
	<li>Point forceps towards the anterior of the fly, grasp it by the thorax, and immerse it in the PBS drop. While viewing through the dissecting microscope, use another pair of forceps to grasp and pull the external genitalia (dark brown structure located at the posterior end of the ventral abdomen) posteriorly until it detaches from the abdomen. In most cases, the testes, seminal vesicles, and accessory gland will be removed from the abdomen along with the external genitalia; if not, insert a single pair of forceps into the abdomen and tease out the testes.</li>
	<li>Separate testes from accessory gland and external genitalia using two pairs of forceps in the PBS drop (Figure 2). Wild-type testes are easily distinguished from neighboring white tissues by their yellow color. Proceed immediately to step 2.</li>
	<li>To isolate testes from pharate males (i.e. enclosed within the pupal case), an additional step must first be performed that involves removing the fly from the pupal case; this step has been previously described elsewhere31. Proceed with dissection as for the adult testes beginning at step 1.2.</li>
	<li>To isolate testes from larval males, perform a modification of a protocol for isolating Drosophila larval ovaries32. Briefly, male larvae can be distinguished from female larvae by the presence of a pair of large, clear, oval structures (larval testes) embedded in the posterior third of the fat body. To isolate larval testes, partially flay open the male larva to isolate the testes and the surrounding fat body from the abdomen as described for the isolation of larval ovaries. Proceed immediately to step 2 of the protocol described herein; the testes can later be removed from the fat body just prior to mounting (steps 2.3 or 3.14) as described for the ovaries32.</li>
</ol>
2.&#160;Sample Preparation and Live Imaging
<ol>
	<li>Use a pair of forceps to gently place 2-3 pairs of testes in a drop of 4-5 ml of PBS on a square glass cover slip. Note that the ratio of testes number to PBS volume may need to be adjusted: too much liquid will prevent cells from spreading properly when squashed, whereas too little liquid will cause cells to burst when squashed. Optional: Use siliconized cover slips to minimize adherence of tissue to cover slip in step 3.2.</li>
	<li>Use a pair of forceps to tear open each testis at an appropriate position so as to maximize the presence of the desired germline cell types in the preparation (note that the contents of the testis will mostly egress from the torn region onto the slide during the squashing step.) To enrich for spermatogonia and spermatocytes, tear open the testis adjacent to its apical tip (level 1, Figure 2B). To enrich for spermatocytes and spermatids, tear open the testis at a position slightly basal to level 1 (level 2, Figure 2B). To enrich for more mature germline cells, tear open the testis closer to where the curvature begins (level 3, Figure 2B).</li>
	<li>Gently place a glass microscope slide over the cover slip to squash the testes; do not apply pressure manually as the weight of the cover slip alone is sufficient to obtain a properly squashed sample. Try to avoid trapping air bubbles. Optional: Use poly-L-lysine coated microscope slides to promote adherence of tissue to slide in step 3.2.</li>
	<li>Use preparation immediately (ideally within 15 min of preparation) to observe live cells by phase-contrast microscopy; for transgenic flies with expression of fluorescently tagged proteins in the testes, live cells can be examined by fluorescence microscopy at this step. Alternatively, proceed with fixation and antibody staining (Protocol 3).</li>
	<li>Gently wick any excess liquid from under the coverslip using a cleaning wipe to allow flattening of the preparation until the germ cells are clearly in focus.</li>
</ol>
3.&#160;Formaldehyde Fixation and Antibody Staining
<ol>
	<li>Snap freeze each slide containing squashed testes (from Protocol 2) using a pair of metal tongs to immerse it briefly in liquid nitrogen (until liquid nitrogen stops bubbling).</li>
	<li>Remove the cover slip immediately using a razor blade.</li>
	<li>Use metal tongs to transfer slides to a prechilled glass slide rack filled with ice-cold 95% ethanol (spectrophotometric grade, methanol-free). Store at -20 &#176;C for 10 min.</li>
	<li>Use metal tongs to transfer slides to a glass slide rack filled with 4% formaldehyde in PBS plus 0.1% Triton X-100 (PBS-T). Store at room temperature for 7 min.</li>
	<li>Use metal tongs to transfer slides to a glass slide rack filled with PBS. Wash slides in PBS for 5 min at room temperature. Repeat 1x. Perform all washes by discarding solution in the glass slide rack (i.e. by pouring it out) and replacing with fresh solution.</li>
	<li>Discard the PBS and immerse slides in PBS-T for 30 min at room temperature to permeabilize cell membranes.</li>
	<li>Wash slides in PBS for 5 min at room temperature. Repeat 2x.</li>
	<li>Blocking step (optional): Immerse slides in PBS plus 1% BSA for 45 min at room temperature.</li>
	<li>Use a hydrophobic barrier pen to draw a circle on the slide around squashed tissue (easily visible by eye) in order to confine the antibody solutions (added in steps 3.9 and 3.11). The tissue should be kept moist at all times while performing immunostaining.</li>
	<li>Add 30-40 ml of primary antibody (diluted in PBS-T, 1:400 to 1:50, depending on antibody) to tissue within the circle. If blocking was performed, dilute primary antibody in PBS-T plus 1% BSA. Anti-gamma-tubulin antibody (for staining of centrosomes in Figure 4) was diluted 1:100. Incubate in a moist, dark chamber (e.g. closed plastic box with damp paper towels) for 2 hr at room temperature or overnight at 4 &#176;C.</li>
	<li>Wash slides in PBS for 5 min at room temperature 3x. If blocking was performed, wash twice in PBS-T and once in PBS (5 min at room temperature each).</li>
	<li>Add 30-40 ml of fluorophore-conjugated secondary antibody (diluted 1:400 in PBS) to the tissue and incubate in the dark for 1-2 hr at room temperature.</li>
	<li>Wash slides in PBS for 5 min at room temperature. Repeat 2x.</li>
	<li>Add 30-40 ml of DAPI solution (0.2 mg/ml in PBS) to the tissue within the circle.</li>
	<li>Gently place a glass cover slip over the tissue, taking care to avoid trapping air bubbles. If air bubbles should appear, carefully move around the cover slip without destroying the squash until the bubbles escape from the sides of the cover slip.</li>
	<li>Use a cleaning wipe to blot excess DAPI from the edges of the slide.</li>
	<li>Seal the cover slip to the slide using clear nail polish.</li>
	<li>Use this preparation within the next 3-4 hr to view immunostained cells by fluorescent microscopy. For longer-term storage of slides (up to at least several weeks), use a glycerol-based hard mount media with DAPI, and store slides at -20 &#730;C.</li>
	<li>Alternatively, fix samples in methanol instead of formaldehyde (depending on the antigen). After completing the entire protocol through step 3.2, immerse slides of squashed testes in methanol for 10 min at -20 &#176;C and proceed with step 3.5 onward.</li>
</ol>

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The authors declare that they have no competing financial interests.

Although the testes of wild-type flies can be readily identified due to their yellow color (in contrast the neighboring white tissues), the testes of white mutant flies are white and thus can occasionally be confused with the gut. Most transgenic strains, which are typically in a white background, also have white testes because the mini-white gene found in P-elements does not promote pigment accumulation in the testes. When Drosophila testes cannot be distinguished by color, o...

Drosophila testes are an ideal model system for the study of many biological processes including the regulation of stem cells, meiosis, and sperm development13-18. The spermatocytes and their meiotic spindles are large and hence convenient for cytological analysis, and relaxed cell cycle checkpoints during spermatogenesis facilitate the study of mutations in cell cycle genes. Different cell types can be observed in ordered progression along the length of the testes, and any disruption in spermatogenesis can lead to changes in this overall arrangement. These features combined with Drosophila genetic tools have facilitated the mutational analysis of spermatogenesis21-23.
The stages of Drosophila spermatogenesis have been well defined. Germline cells that develop synchronously within cysts progress sequentially through the stages of spermatogenesis along the length of the testis. During both the mitotic and meiotic divisions of the male germ cells, cytokinesis occurs incompletely such that the daughter cells remain connected by cytoplasmic bridges known as ring canals (Figure 1). The apical tip of the testis contains a population of germline stem cells that gives rise to spermatogonial cells, which undergo four mitotic divisions with incomplete cytokinesis to generate 16-cell cysts of primary spermatocytes. After premeiotic S phase, primary spermatocytes enter G2, a prolonged growth period of ~90 hr during which cellular volume increases ~25-fold. Progression through meiosis I and meiosis II results in the formation of 32-cell cysts of secondary spermatocytes and 64-cell cysts of haploid spermatids, respectively. The immature, round spermatids undergo extensive cellular remodeling to form mature sperm. Post-meiotic cells, in particular the bundles of elongating and mature spermatids, occupy much of the volume of the testis.
The successful transport of functional sperm to female flies requires coordination between the different parts of the male reproductive system, which is composed of several paired structures (the testes, seminal vesicles, and accessory glands) and a single ejaculatory duct (Figure 2). Sperm are produced within the testes and stored within the seminal vesicles until copulation24. The accessory glands contain secretory cells that produce seminal fluid. The sperm migrating from the seminal vesicles are mixed with seminal fluid within the ejaculatory duct, which is connected to both the seminal vesicles and the accessory glands. This mixture of sperm and seminal fluid is ultimately pumped out of the male into the vagina of the female fly through the ejaculatory bulb located at the posterior end of the male abdomen25. Proteins within the seminal fluid are essential for prolonged storage of sperm within specialized organs known as spermathecae in the reproductive tract of Drosophila females26.
Excellent methods for the isolation of Drosophila testes and visualization of cells at various stages of spermatogenesis are available in the scientific literature3-12. We herein add to this body of knowledge by presenting examples of these protocols with an accompanying video demonstration. The protocol for preparation of live testes samples for phase-contrast microscopy is based on a previously described method27. The protocol for formaldehyde fixation and immunostaining of testes is also based on a previously described method28. The approaches described herein have been used in many studies of Drosophila spermatogenesis (for example, to assess the roles of dynein, a minus-end-directed microtubule motor, during Drosophila spermatogenesis).
In addition to the basic protocols, suggestions are provided for varying the dissection so as to enrich for spermatogonia, spermatocytes, or mature sperm. Different methods for processing the testes such that cysts either remain intact or are disrupted as needed are described. An advantage in using Drosophila testes as a model system is that, compared to Drosophila oocytes and embryos, antibodies and dyes can easily penetrate cells following their dispersal from the testes, and fewer washing steps are required; thus, protocols can be performed in a relatively short time.

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cytological analysis of spermatogenesis: live and fixed preparations of drosophila testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both the female and male germ lines of Drosophila have contributed greatly to the current understanding of meiosis as well as stem cell biology. Excellent protocols are available in the literature for the isolation and imaging of Drosophila ovaries&#160;and testes3-12. Herein, methods for the dissection and preparation of Drosophila testes for microscopic analysis are described with an accompanying video demonstration. A protocol for isolating testes from the abdomen of adult males and preparing slides of live tissue for analysis by phase-contrast microscopy as well as a protocol for fixing and immunostaining testes for analysis by fluorescence microscopy are presented. These techniques can be applied in the characterization of Drosophila mutants that exhibit defects in spermatogenesis as well as in the visualization of subcellular localizations of proteins.

An example of a properly dissected pair of Drosophila male reproductive organs is shown in Figure 2A. Testes removed from the abdomen of the adult male fly are typically attached to the ejaculatory duct (brown, Figure 2A&#39;) and a pair of accessory glands (green, Figure 2A&#39;) via a pair of seminal vesicles (blue, Figure 2A&#39;). To separate the testes from most of the accompanying somatic tissue, the ejaculatory duct and the accessory glan...

Watch this Scientific Journal Video about Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes at JoVE.com

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Methods for isolating and preparing Drosophila testes samples (live and fixed) for imaging by phase-contrast and fluorescence microscopy are described herein.&#160;

Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Drosophila melanogaster is a powerful model system that has been widely used to elucidate a variety of biological processes. For example, studies of both ...

cytological-analysis-spermatogenesis-live-fixed-preparations

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26.0K vistas.  Vanderbilt University Medical Center.  Los métodos para aislar y preparar muestras de Drosophila testículos (en directo y fijo) para obtener imágenes de contraste de fase y microscopía de fluorescencia se describen en este documento. 

Video: Cytological Analysis of Spermatogenesis: Live and Fixed Preparations of Drosophila Testes

Thin Sectioning of Slice Preparations for Immunohistochemistry

Many investigations in neuroscience, as well as other disciplines, involve studying small, yet macroscopic pieces or sections of tissue that have been preserved, freshly removed, or excised but kept viable, as in slice preparations of brain tissue.  Subsequent microscopic studies of this material can be challenging, as the tissue samples may be difficult to handle.  Demonstrated here is a method for obtaining thin cryostat sections of tissue with a thickness that may range from 0.2-5.0 mm.  We routinely cut 400 micron thick Vibratome brain slices serially into 5-10 micron coronal cryostat sections.  The slices are typically first used for electrophysiology experiments and then require microscopic analysis of the cytoarchitecture of the region from which the recordings were observed.  We have constructed a simple device that allows controlled and reproducible preparation and positioning of the tissue slice.  This device consists of a cylinder 5 cm in length with a diameter of 1.2 cm, which serves as a freezing stage for the slice.  A ring snugly slides over the cylinder providing walls around the slice allowing the tissue to be immersed in freezing compound (e.g., OCT).  This is then quickly frozen with crushed dry ice and the resulting  wafer  can be position easily for cryostat sectioning.  Thin sections can be thaw-mounted onto coated slides to allow further studies to be performed, such as various staining methods, in situ hybridization, or immunohistochemistry, as demonstrated here.

The present method allows reproducible cryostat sectioning of small, difficult-to-manage, tissue pieces, such as biopsies and brain slices. We utilize a simple aluminum freezing stage to facilitate handling of tissue and a standard cryostat to routinely produce 5-10 micron serial sections from 400 micron thick brain slices.

Many investigations in neuroscience, as well as other disciplines, involve studying small, yet macroscopic pieces or sections of tissue that have been ...

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 ...

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 ...

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 ...

Quantitative Live Cell Fluorescence-microscopy Analysis of Fission Yeast

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using fluorochromes like Green Fluorescent Protein (GFP) directly attached to the protein of interest has become increasingly popular 1. Using GFP and similar fluorochromes the subcellular localisations and movements of proteins can be detected in a fluorescent microscope. Moreover, also the subnuclear localisation of a certain region of a chromosome can be studied using this technique. GFP is fused to the Lac Repressor protein (LacR) and ectopically expressed in the cell where tandem repeats of the lacO sequence has been inserted into the region of interest on the chromosome2. The LacR-GFP will bind to the lacO repeats and that area of the genome will be visible as a green dot in the fluorescence microscope. Yeast is especially suited for this type of manipulation since homologous recombination is very efficient and thereby enables targeted integration of the lacO repeats and engineered fusion proteins with GFP 3. Here we describe a quantitative method for live cell analysis of fission yeast. Additional protocols for live cell analysis of fission yeast can be found, for example on how to make a movie of the meiotic chromosomal behaviour 4. In this particular experiment we focus on subnuclear organisation and how it is affected during gene induction. We have labelled a gene cluster, named Chr1, by the introduction of lacO binding sites in the vicinity of the genes. The gene cluster is enriched for genes that are induced early during nitrogen starvation of fission yeast 5. In the strain the nuclear membrane (NM) is labelled by the attachment of mCherry to the NM protein Cut11 giving rise to a red fluorescent signal. The Spindle Pole body (SPB) compound Sid4 is fused to Red Fluorescent Protein (Sid4-mRFP) 6. In vegetatively growing yeast cells the centromeres are always attached to the SPB that is embedded in the NM 7. The SPB is identified as a large round structure in the NM. By imaging before and 20 minutes after depletion of the nitrogen source we can determine the distance between the gene cluster (GFP) and the NM/SPB. The mean or median distances before and after nitrogen depletion are compared and we can thus quantify whether or not there is a shift in subcellular localisation of the gene cluster after nitrogen depletion.

The fission yeast, Schizosaccharomyces pombe, is a good model system to study basic cellular processes. Here we describe a method to perform quantitative live cell analysis of fission yeast. In this particular experiment we focus on organisation of the genome within the cell nucleus, but the method can also be used to study cytosolic factors.

Several microscopy techniques are available today that can detect a specific protein within the cell. During the last decade live cell imaging using ...

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 ...

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  ...

Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

Flow cytometry has been widely used to obtain information about DNA content in a population of cells, to infer relative percentages in different cell cycle phases. This technique has been successfully extended to the mitotic tissues of the model organism Drosophila melanogaster for genetic studies of cell cycle regulation in vivo. When coupled with cell-type specific fluorescent protein expression and genetic manipulations, one can obtain detailed information about effects on cell number, cell size and cell cycle phasing in vivo. However this live-cell method has relied on the use of the cell permeable Hoechst 33342 DNA-intercalating dye, limiting users to flow cytometers equipped with a UV laser. We have modified this protocol to use a newer live-cell DNA dye, Vybrant DyeCycle Violet, compatible with the more common violet 405nm laser. The protocol presented here allows for efficient cell cycle analysis coupled with cell type, relative cell size and cell number information, in a variety of Drosophila tissues. This protocol extends the useful cell cycle analysis technique for live Drosophila tissues to a small benchtop analyzer, the Attune Acoustic Focusing Cytometer, which can be run and maintained on a single-lab scale.

Live Cell Cycle Analysis of Drosophila Tissues using the Attune Acoustic Focusing Cytometer and Vybrant DyeCycle Violet DNA Stain

A protocol for cell cycle analysis of live Drosophila tissues using the Attune Acoustic Focusing Cytometer is described. This protocol simultaneously provides information about relative cell size, cell number, DNA content and cell type via lineage tracing or tissue specific expression of fluorescent proteins in vivo.

Flow cytometry has  been widely used to obtain information about DNA content in a population of  cells, to infer relative percentages in different cell ...

Analysis of Embryonic and Larval Zebrafish Skeletal Myofibers from Dissociated Preparations

The zebrafish has proven to be a valuable model system for exploring skeletal muscle function and for studying human muscle diseases. Despite the many advantages offered by in vivo analysis of skeletal muscle in the zebrafish, visualizing the complex and finely structured protein milieu responsible for muscle function, especially in whole embryos, can be problematic. This hindrance stems from the small size of zebrafish skeletal muscle (60 &mu;m) and the even smaller size of the sarcomere. Here we describe and demonstrate a simple and rapid method for isolating skeletal myofibers from zebrafish embryos and larvae. We also include protocols that illustrate post preparation techniques useful for analyzing muscle structure and function. Specifically, we detail the subsequent immunocytochemical localization of skeletal muscle proteins and the qualitative analysis of stimulated calcium release via live cell calcium imaging. Overall, this video article provides a straight-forward and efficient method for the isolation and characterization of zebrafish skeletal myofibers, a technique which provides a conduit for myriad subsequent studies of muscle structure and function.

Zebrafish are an emerging system for modeling human disorders of the skeletal muscle. We describe a fast and efficient method to isolate skeletal muscle myofibers from embryonic and larval zebrafish. This method yields a high-density myofiber preparation suitable for study of single skeletal muscle fiber morphology, protein subcellular localization, and muscle physiology.

The zebrafish has  proven to be a valuable model system for exploring skeletal muscle function and  for studying human muscle diseases. Despite  the many ...