This work was supported by a grant from the National Science Foundation.

1. Day 1
Before you start: 
A healthy population of flies should be maintained using standard culturing protocols: remove adults from bottles or vials after 3-4 days of egg-lay and allow development to proceed at a constant temperature until wandering 3rd instar larvae initiate pupariation. The larval/pupal transition is marked by the formation of the prepupae (considered 0 hours after puparium formation-APF). Immobile pupae are distinguished from older pupae by their white coloration and from larvae that have not yet begun pupariation by their oblong,rounded shape and protrusion of the anterior spiracles. 
You will need:
<ul>
<li>A paint brush for collecting pupae</li>
<li>A humid chamber for culturing pupae: A Petri dish lined with wetted paper towel and covered with filter paper marked for the various time points of collection, genotype or sex of the pupae</li>
<li>1X Phosphate buffered saline (PBS) for washing pupae</li>
</ul>
Collection, culturing and staging pupae
<ol>
<li>Using a wetted paintbrush gently remove 0hr APF pupae from culture bottles/vials and place them in the lid of the humid chamber. </li>
<li>Using the paintbrush and 1X PBS gently wash the pupae to remove debris from the pupa case</li>
<li>If necessary sort the pupae by sex. The gonad primordia of pupae can be used to sort the sexes. The male gonad primordia are a lateral pair of translucent discs easily seen through the pupal cuticle approximately 2/3 down the length of the body 5. The female gonad primordia are smaller and not as easily identified.</li>
<li>Place the pupae in the appropriately labeled position in the humid chamber and return to constant temperature. Culture to the appropriate developmental time point.</li>
</ol>
2. Day 2: Dissection, fixation and primary antibody incubation
Before you start:
You will need:
<ul>
<li>Dissection stereomicroscope</li>
<li>Surgical forceps</li>
<li>Surgical scalpel with number 11 blades</li>
<li>Two nine-well glass depression dishes (Corning product # 7220-85)<ul>
<li>Fill wells of one dish with 1X PBS: This is the rinsing dish for cleaning dissected pupae</li>
<li>Fill wells of second dish with fixation buffer</li></ul></li>
<li>Humid chamber for antibody incubation. We use plastic sealable sandwich boxes lined with wetted paper towels.</li>
<li>Fixation buffer: 1x PBS, 4% paraformaldehyde, 0.2% deoxycholic acid (for permeabilization)</li>
<li>Dissection platform: Clear microscope slide with a piece of double-sided tape adhered to one side</li>
<li>1X PBS</li>
<li>p100 or p200 pipettor</li>
</ul>
Dissection
<ol>
<li>Using forceps gently remove pupae one at a time and place them on the dissection platform with the anterior end of the pupae facing the broadest width of the tape. If pupae are excessively wet, first blot them dry using a paper towel.</li>
<li>Before the pupae have completely adhered to the tape, use a paintbrush to position them.</li>
<li>Allow the pupae to air dry and adhere to the tape (5-10 minutes)</li>
<li>With the surgical scalpel dissect each pupae bilaterally. This is best accomplished with a single rapid cut from the anterior to the posterior end of the pupae. If properly done, the cut will bisect both anterior and posterior spiracles pairs. Dissect no more than 10 pupae at a time as washing and cleaning the samples quickly is necessary to prevent proteolytic damage to the tissue.</li>
<li>Using a paintbrush, transfer a small amount of 1X PBS to each dissected pupae to loosen them from the tape.</li>
</ol>
Cleaning, fixation and primary antibody incubation 
<ol start="6">
<li>With a pair of surgical forceps grasp an individual pupa half anteriorly and immediately immerse it in a well of the rinsing dish. The dissection platform should be replaced by the rinsing dish on the microscope stage. The pupae case should be left on the sample until processing is complete and samples are ready for mounting. </li>
<li>While still grasping the pupa half use a pipettor to gently wash away the internal tissue of the abdomen. Too much pressure during washing can lead to loss of epithelial cells.</li>
<li>Immediately transfer the clean pupa half to fixation buffer at room temperature and process the remaining pupa halves similarly. With practice, dissection and washing of 20 pupa halves should take less than 5 minutes.</li>
<li>Allow pupae to incubate in fixation buffer at room temperature for 1 (one) hour.</li>
<li>Rinse fixed pupae 3x five minutes in 1X PBS</li>
<li>Samples may immediately be processed for immunohistochemistry or may be stored in 100% ethanol at -20&deg;C for up to 3 months without loss of cells or epitope reactivity.</li>
<li>To store samples, equilibrate samples through a dilution series of PBS:EtOH (3:1, 1:1, 1:3) at room temperature for 20 minutes in each dilution prior to transferring to 100% EtOH.</li>
<li>Samples must be re-equilibrated in PBS through the reverse series before processing for immunohistochemistry.</li>
<li>Block samples in 1x PBS (supplemented with 2% bovine serum albumin) for 1 (one) hour before addition of primary antibody.</li>
<li>Incubate with primary antibody diluted to appropriate concentration in 1X PBS over night at 4&deg;C without rocking.</li>
</ol>
3. Day 3: Secondary antibody incubation, mounting and imaging
Before you start:
You will need:
<ul>
<li>Two surgical forceps</li>
<li>Mounting media (glycerol, Vectashield [Vector Labs], or Slowfade Gold [Invitrogen])</li>
<li>Depression well microscope slides (0.8 mm)</li>
<li>Coverslips (24 x 40 mm)</li>
</ul>
Secondary antibody incubation and mounting:
<ol>
<li>Wash samples 3x 10 minute each with 1x PBS</li>
<li>Block samples in 1x PBS (supplemented with 2% bovine serum albumin) for 1 (one) hour before addition of secondary antibody.</li>
<li>Incubate with secondary antibody diluted to appropriate concentration in 1X PBS at room temperature in the dark without rocking for 3 (three) hours.</li>
<li>Wash samples 3x 10 minute each with 1x PBS</li>
<li>If appropriate, counterstain pupae (Example we stain nuclei with DAPI [4',6-diamidino-2-phenylindole] diluted in PBS for 10 minutes).</li>
<li>Equilibrate samples (minimum of 30 minutes) in appropriate media before mounting.</li>
<li>Prepare a sample slide. The morphology of the pupae prevents flat mounting. Samples are mounted in the well of a depression slide (drop slide) for imaging. Place 75-100ul of mounting media in the depression well. Several samples can be mounted on a single slide.</li>
<li>The pupa case must be removed from the samples before mounting. Grasp an individual pupa case anteriorly being sure not to grasp the internal pupal membrane.</li>
<li>With a second pair of forceps gently grasp the internal pupal membrane by the head and remove it from the pupal case.</li>
<li>Immediately transfer the sample to the depression well and continue processing additional samples.</li>
<li>Using a probe or forceps position the samples uniformly in the depression well with the lateral surface facing up. Occasional air bubbles can be removed with a probe.</li>
<li>Gently lower the coverslip onto the samples taking care not to introduce air bubbles</li>
</ol>
4. Representative results:
Samples prepared using this protocol retain the gross morphology of the adult abdomen. Image stacks may be projected to generate a two-dimensional image or 3-D rendering may be applied to investigate abdomen topology.
<img src="/files/ftp_upload/3139/3139fig1.jpg" alt="Figure 1"/> Figure 1. Segmentation gene products in Drosophila pupae. Wingless protein and Engrailed expression (En-gal4:uas-GFP) were visualized at 26 hour after puparium formation (APF) with mouse anti-Wg (4D4: Iowa Hybridoma Bank) and anti-GFP. 10x magnification, anterior is left and dorsal is up. Nucleii were counterstained with DAPI. 
Stacks of approximately 50 images (&Delta;Z between slices is 2.5&mu;m) were projected.

<ol>
	<li>Kopp, A., Duncan, I. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anteroposterior+patterning+in+adult+abdominal+segments+of+Drosophila.">Anteroposterior patterning in adult abdominal segments of Drosophila.</a> Dev. Biol. 242, 15-30 (2002).</li><li>Ninov, N., Chiarelli, D. A., Martin-Blanco, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Extrinsic+and+intrinsic+mechanisms+directing+epithelial+cell+sheet+replacement+during+Drosophila+metamorphosis.">Extrinsic and intrinsic mechanisms directing epithelial cell sheet replacement during Drosophila metamorphosis.</a> Development. 134, 367-379 (2007).</li><li>Bischoff, M., Cseresnyes, Z. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+rearrangements,+cell+divisions+and+cell+death+in+a+migrating+epithelial+sheet+in+the+abdomen+of+Drosophila.">Cell rearrangements, cell divisions and cell death in a migrating epithelial sheet in the abdomen of Drosophila.</a> Development. 136, 2403-2411 (2009).</li><li>Matsuda, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=in+Morphology+and+evolution+of+the+insect+abdomen.">in Morphology and evolution of the insect abdomen.</a> , (1976).</li><li>Kerkis, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Growth+of+the+Gonads+in+Drosophila+Melanogaster.">The Growth of the Gonads in Drosophila Melanogaster.</a> Genetics. 16, 212-224 (1931).</li></ol>

No conflicts of interest declared.

The techniques presented in this video can be used to prepare Drosophila pupae from a variety of developmental time points. Pupae processed during the period of 24 hours APF to 32 hours APF are most prone to cell loss from the epithelium. The use of detergents (such as Triton X-100 and Tween-20) during extended incubation steps increases the likelihood of cell loss and is therefore not recommended. Rather, deoxycholic acid is used as a detergent during the initial fixation step. All subsequent steps are performed in 1X PBS without detergent. Additionally, rocking samples during extended incubation steps increases cell loss and should be avoided.
Samples can be imaged using confocal microscopy techniques. However, Drosophila pupae processed as described can also be imaged using a structured illumination microscopy system yielding image quality comparable to confocal techniques.

<ul>
	<li>Dissection stereomicroscope</li>
	<li>Surgical forceps</li>
	<li>Surgical scalpel with number 11 blades</li>
	<li>Nine-well glass depression dishes (Corning product # 7220-85)</li>
	<li>Humid chamber for culturing pupae</li>
	<li>1X Phosphate buffered saline (PBS)</li>
	<li>#11 surgical scalpel</li>
	<li>double-sided tape</li>
	<li>Fixation buffer: 1x PBS, 4% paraformaldehyde, 0.2% deoxycholic acid (for permeabilization)</li>
	<li>p100 or p200 pipettor</li>
	<li>Depression well microscope slides (0.8 mm)</li>
</ul>

drosophila pupal abdomen immunohistochemistry

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic morphologies 1-3. During pupation, which spans approximately 96 hours (at 25 &deg;C), proliferating populations of imaginal cells replace the larval epidermis to generate the adult abdominal segments. These imaginal cells, born during embryogenesis, exist as lateral pairs of histoblast nests in each abdominal segment of the larvae. Four pairs of histoblast nests give rise to the adult dorsal cuticle (anterior and posterior dorsal nests), the ventral cuticle (ventral nests) and the spiracles associated with each segment (spiracle nests) 4. Upon puparation, these diploid cells (distinguishable by size from the larger polyploid larval epidermal cells- LECs) begin a stereotypical process of proliferation, migration and replacement of the LECs. Various molecular and genetic tools can be employed to investigate the contributions of genetic pathways involved in morphogenesis of the adult abdomen. Ultimate adult phenotypes are typically analyzed following dissection of adult abdominal cuticles. However, investigation of the underlying molecular processes requires immunohistochemical analyses of the pupal epithelium, which present unique challenges. Temporally dynamic morphogenesis and the interactions of two distinct epithelial populations (larval and imaginal) generate a fragile tissue prone to excessive cell loss during dissection and subsequent processing. We have developed methods of dissection, fixation, mounting and imaging of the Drosophila pupal abdominem epithelium for immunohistochemical studies that generate consistent high quality samples suitable for confocal or standard fluorescent microscopy.

Watch this Scientific Journal Video about Drosophila Pupal Abdomen Immunohistochemistry at JoVE.com

Drosophila Pupal Abdomen Immunohistochemistry

Antibody staining of the Drosophila pupae can enhance genetic analyses of adult abdominal developmental genetics. We present our protocol for dissection, fixation and antibody staining of staged Drosophila pupal abdomen.

Drosophila Pupal Abdomen Immunohistochemistry

The Drosophila pupal abdomen is an established model system for the study of epithelial morphogenesis and the development of sexually dimorphic ...

drosophila-pupal-abdomen-immunohistochemistry

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16.0K ビュー.  University of Alabama. の抗体染色ショウジョウバエ蛹は成人腹部発生遺伝学の遺伝学的解析を強化することができます。我々は段階の解剖、固定し、抗体染色のために我々のプロトコルを提示ショウジョウバエ蛹の腹部。

ビデオ: Drosophila Pupal Abdomen Immunohistochemistry

Antibody staining of the Drosophila pupae can enhance genetic analyses of adult abdominal developmental genetics. We present our protocol for dissection, fixation and antibody staining of staged Drosophila pupal...

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Dissection, Immunohistochemistry and Mounting of Larval and Adult Drosophila Brains for Optic Lobe Visualization

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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 ...

In situ Protocol for Butterfly Pupal Wings Using Riboprobes

Here we present, in video format, a protocol for in situ hybridizations in pupal wings of the butterfly Bicyclus anynana using riboprobes.  In situ hybridizations, a mainstay of developmental biology, are useful to study the spatial and temporal patterns of gene expression in developing tissues at the level of transcription. If antibodies that target the protein products of gene transcription have not yet been developed, and/or there are multiple gene copies of a particular protein in the genome that cannot be differentiated using available antibodies, in situs can be used instead. While an in situ technique for larval wing discs has been available to the butterfly community for several years, the current protocol has been optimized for the larger and more fragile pupal wings.

In order to examine gene expression in the pupal wing tissue of Bicyclus anynana, we present an optimized protocol for in situ hybridizations using riboprobes. We also provide guidelines for the further optimization of this protocol for use in pupal wings of other Lepidopteran species.

Here we present, in video format, a protocol for in situ hybridizations in pupal wings of the butterfly Bicyclus anynana using riboprobes.  In situ ...

Immunohistochemistry: Paraffin Sections Using the Vectastain ABC Kit from Vector Labs

Immunohistochemistry (IHC) is a valuable technique utilized to localize/visualize protein expression in a mounted tissue section using specific antibodies. There are two methods: the direct and indirect method. In this experiment, we will only describe the use of indirect IHC staining. Indirect IHC staining utilizes highly specific primary and biotin-conjugated secondary antibodies. Primary antibodies are utilized to discretely identify proteins of interest by binding to a specific epitope, while secondary antibodies subtract for non-specific background staining and amplify signal by forming complexes to the primary antibody. Slides can either be generated from frozen sections, or paraffin embedded sections mounted on glass slides. In this protocol, we discuss the preparation of paraffin-embedded sections by dewaxing, hydration using an alcohol gradient, heat induced antigen retrieval, and blocking of endogenous peroxidase activity and non-specific binding sites. Some sections are then stained with antibodies specific for T cell marker CD8 and while others are stained for tyrosine hydroxylase. The slides are subsequently treated with appropriate secondary antibodies conjugated to biotin, then developed utilizing avidin-conjugated horseradish peroxidase (HRP) with Diaminiobenzidine (DAB) as substrate. Following development, the slides are counterstained for contrast, and mounted under coverslips with permount. After adequate drying, these slides are then ready for imaging.

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Immunohistochemistry on Paraffin Sections of Mouse Epidermis Using Fluorescent Antibodies

In the epidermis, immunohistochemistry is an efficient means of localizing specific proteins to their relative expression compartment; namely the basal, suprabasal, and stratum corneum layers. The precise localization within the epidermis of a particular protein lends clues toward its functional role within the epidermis. In this chapter, we describe a reliable method for immunolocalization within the epidermis modified for both frozen and paraffin sections that we use very routinely in our laboratory. Paraffin sections generally provide much better morphology, hence, superior results and photographs; however, not all antibodies will work with the harsh fixation and treatment involved in their processing. Therefore, the protocol for frozen sectioning is also included. Within paraffin sectioning, two fixation protocols are described (Bouin's and paraformaldehyde); the choice of fixative will be directly related to the antibody specifications and may require another fixing method.

We describe a reliable method for immunolocalization within the epidermis modified for both frozen and paraffin sections that we use very routinely in our laboratory.

In the epidermis, immunohistochemistry is an efficient means of localizing specific proteins to their relative expression compartment; namely the basal, ...

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use of the Drosophila motor system is due to its high accessibility. It can be analyzed with single-cell resolution. There are 30 muscles per hemisegment whose arrangement within the peripheral body wall are known. A total of 31 motor neurons attach to these muscles in a pattern that has high fidelity. Using molecular biology and genetics, one can create transgenic animals or mutants. Then, one can study the developmental consequences on the morphology and function of the NMJ. Immunohistochemistry can be used to clearly image the components of the NMJ. In this article, we demonstrate how to use antibody staining to visualize the Drosophila larval NMJ.

This protocol demonstrates how to perform immunohistochemistry on dissected Drosophila larva.

The Drosophila neuromuscular junction (NMJ) is an established model system used for the study of synaptic development and plasticity. The widespread use ...

Production of Tissue Microarrays, Immunohistochemistry Staining and Digitalization Within the Human Protein Atlas

The tissue microarray (TMA) technology provides the means for high-throughput analysis of multiple tissues and cells. The technique is used within the Human Protein Atlas project for global analysis of protein expression patterns in normal human tissues, cancer and cell lines. Here we present the assembly of 1 mm cores, retrieved from microscopically selected representative tissues, into a single recipient TMA block. The number and size of cores in a TMA block can be varied from approximately forty 2 mm cores to hundreds of 0.6 mm cores. The advantage of using TMA technology is that large amount of data can rapidly be obtained using a single immunostaining protocol to avoid experimental variability. Importantly, only limited amount of scarce tissue is needed, which allows for the analysis of large patient cohorts 1 2. Approximately 250 consecutive sections (4 &mu;m thick) can be cut from a TMA block and used for immunohistochemical staining to determine specific protein expression patterns for 250 different antibodies. In the Human Protein Atlas project, antibodies are generated towards all human proteins and used to acquire corresponding protein profiles in both normal human tissues from 144 individuals and cancer tissues from 216 different patients, representing the 20 most common forms of human cancer. Immunohistochemically stained TMA sections on glass slides are scanned to create high-resolution images from which pathologists can interpret and annotate the outcome of immunohistochemistry. Images together with corresponding pathology-based annotation data are made publically available for the research community through the Human Protein Atlas portal (<a href="http://www.proteinatlas.org" target="_blank">www.proteinatlas.org</a>) (Figure 1) 3 4. The Human Protein Atlas provides a map showing the distribution and relative abundance of proteins in the human body. The current version contains over 11 million images with protein expression data for 12.238 unique proteins, corresponding to more than 61% of all proteins encoded by the human genome.

Tissue microarrays allows for an efficient method to gain concurrent information from a multitude of tissues. Representative parts of tissues are assembled into a single paraffin block. Sections from the block are used for immunohistochemistry and analysis of protein expression patterns. Digital scanning generates corresponding images for distribution of data.

The  tissue microarray (TMA) technology provides the means for high-throughput  analysis of multiple tissues and cells. The technique is used within the ...

Optimized Protocol for Retinal Wholemount Preparation for Imaging and Immunohistochemistry

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ring-supported hydrophilized PTFE membrane to provide structural support to both living and fixed tissue during immunohistochemical processing, which allows for the use of a variety of protocols that would otherwise cause damage to the tissue. First, this is demonstrated with bolus loading of fluorescent markers into living retinal tissue. This method allows for quick visualization of targeted structures, while the membrane support maintains tissue integrity during the injection and allows for easy transfer of the preparation for further imaging or processing.
Second, a procedure for antibody staining in tissue fixed with carbodiimide is described. Though paraformaldehyde fixation is more common, carbodiimide fixation provides a superior substrate for the visualization of synaptic proteins. A limitation of carbodiimide is that the resulting fixed tissue is relatively fragile; however, this is overcome with the use of the supporting membrane. Retinal tissue is used to demonstrate these techniques, but they may be applied to any fragile tissue.

The protocol described here is for structural assessment of a wholemount retinal preparation. This includes descriptions of tissue dissection, mounting onto a hydrophilized polytetrafluoroethylene (PTFE) membrane insert, bolus loading with fluorescent markers, and a comparison of fixation with carbodiimide and paraformaldehyde for immunohistochemical analysis of cellular and synaptic components.

Working with delicate tissue can be a complicating factor when performing immunohistochemical assessment. Here, we present a method that utilizes a ...

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.

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 ...

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes differentiation from a stem cell population, mitotic amplification, and meiosis. In addition, post-meiotic germ cells undergo a dramatic morphological reshaping process as well as a global epigenetic reconfiguration of the germ line chromatin&#8212;the histone-to-protamine switch.
Studying the role of a protein in post-meiotic spermatogenesis using mutagenesis or other genetic tools is often impeded by essential embryonic, pre-meiotic, or meiotic functions of the protein under investigation. The post-meiotic phenotype of a mutant of such a protein could be obscured through an earlier developmental block, or the interpretation of the phenotype could be complicated. The model organism Drosophila melanogaster offers a bypass to this problem: intact testes and even cysts of germ cells dissected from early pupae are able to develop ex vivo in culture medium. Making use of such cultures allows microscopic imaging of living germ cells in testes and of germ-line cysts. Importantly, the cultivated testes and germ cells also become accessible to pharmacological inhibitors, thereby permitting manipulation of enzymatic functions during spermatogenesis, including post-meiotic stages.
The protocol presented describes how to dissect and cultivate pupal testes and germ-line cysts. Information on the development of pupal testes and culture conditions are provided alongside microscope imaging data of live testes and germ-line cysts in culture. We also describe a pharmacological assay to study post-meiotic spermatogenesis, exemplified by an assay targeting the histone-to-protamine switch using the histone acetyltransferase inhibitor anacardic acid. In principle, this cultivation method could be adapted to address many other research questions in pre- and post-meiotic spermatogenesis.

This protocol describes the dissection and cultivation of intact testes and germ-line cysts from Drosophila melanogaster pupae. This method allows microscopic observation of spermatogenesis ex vivo. Furthermore, we describe a pharmacological assay of the effect of inhibitors on specific stages of germ-cell development in pupal testes.

During spermatogenesis in mammals and in Drosophila melanogaster, male germ cells develop in a series of essential developmental processes. This includes ...

The Drosophila melanogaster eye disc is a powerful system that can be used to study many different biological processes. It contains approximately 800 separate eye units, termed ommatidia1. Each ommatidium contains eight neuronal photoreceptors that develop from undifferentiated cells following the passage of the morphogenetic furrow in the third larval instar2. Following the sequential differentiation of the photoreceptors, non-neuronal cells develop, including cone and pigment cells, along with mechanosensory bristle cells3. Final differentiation processes, including the structured arrangement of all the ommatidial cell types, programmed cell death of undifferentiated cell types and rhodopsin expression, occurs through the pupal phase4-7. This technique focuses on manipulating the pupal eye disc, providing insight and instruction on how to dissect the eye disc during the pupal phase, which is inherently more difficult to perform than the commonly dissected third instar eye disc. This technique also provides details on immunostaining to allow the visualization of various proteins and other cell components.

The goal of this technique is to enable researchers to perform dissection, immunostaining and mounting of pupal eye discs from Drosophila melanogaster of any age.

The Drosophila melanogaster eye disc is a powerful system that can be used to study many different biological processes.  It contains approximately 800 ...