We thank Marco Antonio Rosales Vega and Abel Segura for taking some of the confocal images, Carmen Mu&#241;oz for media preparation and Dr. Arturo Pimentel, M.C. Andr&#233;s Saralegui, and Dr. Chris Wood from the LMNA for advice on the use of the microscopes.
FUNDING: 
This work was supported by the Consejo Nacional de Ciencia y Tecnolog&#237;a (CONACyT) (A1-S-8239 to VV-G) and Programa de Apoyo a Proyectos de Investigaci&#243;n e Innovaci&#243;n Tecnol&#243;gica (204915 and 200118 to VV-G)

1. Third instar larvae culture
<ol>
	<li>Prepare 1 liter of standard media by adding 100 g of yeast, 100 g of unrefined whole cane sugar, 16 g of agar, 10 mL of propionic acid and 14 g of gelatin. Dissolve all ingredients except the yeast in 800 mL of tap water and then dissolve the yeast. Autoclave immediately for 30 minutes.
	<ol>
		<li>Afterward, let the media cool down to 60 &#176;C and add propionic acid to a final concentration 0.01%. Let the bottle stand until the gelatin is formed.</li>
	</ol>
	</li>
	<li>To optimize the 3&#7506; instar larvae culture, first collect 5-to-10-day old adults and place 50 (25 males and 25 females) in a broad neck bottle of standard media.</li>
	<li>Place the bottle with the flies in a controlled temperature incubator at 25 &#176;C until the number of eggs laid is 50 (approximately 12 hours for the wild-type strain).</li>
	<li>After the incubation time is over, remove the adults and transfer them to a new bottle to repeat the procedure. Let the embryos grow at 18 &#176;C for 72 hours 
	&#8203;NOTE: For more about Drosophila stock maintenance conditions, see Tennessen &#38; Thymmel13.</li>
</ol>
2. Larvae collection
<ol>
	<li>For larvae collection choose the wandering larvae that do not have everted spiracles. After the eversion of the spiracles, the larva enters the prepupal stage, while retaining excellent polytene chromosomes suitable for analysis. Only after 12 hours do the cells of the salivary gland begin to prepare for programmed cell death14,15.</li>
	<li>Take fifteen 3&#176; instar larvae and put them in a watch glass to wash them. Then transfer them to an ice-cold saline solution or PBS (1x PBS: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 2 mM KH2PO4, adjust pH to 7.4).</li>
	<li>Dissect 15 to 30 pairs of salivary glands (or as many as possible in 30 minutes) in cold PBS with protease inhibitors under the stereoscopic microscope. Transfer the salivary glands to a 1.5 mL tube with ice-cold PBS.</li>
	<li>Wash once with 1 mL of PBS plus protease inhibitors. Wait for the tissue to reach the bottom of the tube.</li>
	<li>After the wash, remove the PBS with a 1000 &#181;L pipette. Take care not to touch the tissue.
	<ol>
		<li>Alternatively, dissect the salivary glands in 5 mL of PBS to eliminate the need for this washing step and proceed to step 3 by transferring the salivary glands to 0.5 mL of the Ruvkun fixing buffer described below.</li>
	</ol>
	</li>
</ol>
3. Salivary gland tissue fixation
<ol>
	<li>After removing the PBS from the last step, directly add 0.5 mL of 1x Ruvkun fixing buffer, with 50% methanol (add 0.5 mL of methanol) and 2% formaldehyde. 
	&#8203;NOTE: 2x Ruvkun solution is 160 mM KCl, 40 mM NaCl, 20 mM EGTA, 30 mM PIPES at pH 7.4.</li>
	<li>Incubate for 2 hours at 4 &#176;C with mild rotation.</li>
</ol>
4. Salivary gland tissue wash
<ol>
	<li>Carry out one 5-minute rotation wash with 1 mL of Tris/Triton buffer (100 mM Tris pH 7.4, 1% Triton X-100 and 1 mM EDTA). 
	&#8203;NOTE: Wait for the tissue to reach the bottom of the tube.</li>
</ol>
5. Permeabilization
<ol>
	<li>Incubate the salivary glands in 1 mL of Tris/Triton X-100 (the same as above). For some proteins it might be necessary to add 1% &#946;-mercaptoethanol.</li>
	<li>Incubate for 2 hours at 37 &#176; C with mild shaking (300 rpm).</li>
</ol>
6. Preservation step (optional)
NOTE: If not proceeding immediately to the incubation with the antibody, preserve the tissue as follows.
<ol>
	<li>Wash with 1 mL of BO3 buffer (0.01 M H3BO3 pH 9.2 + 0.01 M NaOH) and then incubate in BO3/10 mM DTT at 37 &#176; C with mild shaking (300 rpm) for 15 minutes.</li>
	<li>At the end of the incubation period, perform a wash with 1 mL of BO3 buffer alone. 
	NOTE: Wait for the tissue to reach the bottom of the tube.</li>
	<li>Add 1 mL of PBS. Preserve the tissue in this solution at 4 &#176;C for up to 72 hours and then proceed with the next step. This step is particularly helpful when working with different mutant strains that may present a delayed life cycle, so the immunodetection can be performed at the same time along with the controls.</li>
</ol>
7. Tissue blocking
<ol>
	<li>Incubate the salivary glands in 1 mL of Buffer B (PBS + 0.1% BSA + 0.5% Triton X-100 + 1 mM EDTA) for 2 hours at room temperature with rotation.</li>
</ol>
8. Immunostaining
<ol>
	<li>Remove all buffer B and add buffer A (PBS + 0.1% BSA) plus antibody of interest. 
	&#8203;NOTE: We use the HP1a C1A9c (concentrated antibody) from Hybridoma Bank up to 1:3000. When using the C1A9s (supernatant) we have tried from 1:100 to a 1:500 dilution and any dilution between this rank works well) overnight at 4 &#7506;C with rotation. At this point it is important that the shaking does not raise bubbles which might damage the antibody.</li>
</ol>
9. Immunostaining washing
<ol>
	<li>Perform 3 x 15-minute washes with buffer B under stirring at room temperature using 1 mL each time.</li>
	<li>Transfer the glands to buffer B together with the secondary antibody coupled to a fluorophore for 2 hours under rotation at 4 &#7506;C (secondary antibody Alexa fluor 568 Invitrogen were used 1:3000).
	<ol>
		<li>Cover the tube with aluminum paper foil to protect the secondary antibody from the light.</li>
	</ol>
	</li>
	<li>Carry out 2 x 15-minute washes at room temperature while rotation with 1 mL of Buffer B.</li>
	<li>Incubate with a DNA marker such as Sytox (take 2 &#181;L of 5 mM stock and dissolve in 1 mL of Buffer B) or Hoechst (take 1 &#181;L of 10 mg/mL stock and dissolve in 1 mL of Buffer B) for 10 minutes at room temperature with rotation.</li>
	<li>Carry out one wash with Buffer B and once with PBS, each wash lasting 10 minutes while rotating at room temperature. 
	&#8203;NOTE: Remember to protect it from the light.</li>
</ol>
10. Imaging
<ol>
	<li>Mount the salivary glands on a slide, making a pool with a coverslip.</li>
	<li>Put the salivary glands in the middle of the pool and cover with AF1 citifluor to avoid the formation of bubbles extending the viscous liquid all over the place. Then seal all the sides with clear nail polish.</li>
	<li>Observe under a fluorescence or confocal microscope. If the sample is not going to be observed on the same day, store away from light at 4 &#176;C.</li>
	<li>Use GraphPad Prism 6 to generate all graphs and statistical analyses.</li>
	<li>Analyze the data from HP1a distribution in salivary glands using the Kruskal-Wallis test. Statistical significance was set at (p &#60; 0.05*, &#60; 0.01 **, &#60; 0.001***, &#60; 0.0001****).</li>
</ol>

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The authors declare that they have no competing interests.

The cellular function of eukaryotic organisms can define the 3D structure within the nucleus, which is supported by interactions between different proteins with chromatin and various molecules including RNA. In the last three years, the biological condensates that have had relevance, including heterochromatin, have taken a fundamental role in the determination of the phase separation promoting the distinct nuclear spatial organization of active and repressive chromatin 16,<sup class="xr...

Since the early studies of Emil Heitz1, heterochromatin has been considered an important regulator of cellular processes such as gene expression, meiotic and mitotic separation of chromosomes, and the maintenance of genome stability2,3,4.
Heterochromatin is mainly divided into two types: constitutive heterochromatin that characteristically defines repetitive sequences, and transposable elements that are present at specific chromosome sites such as the telomeres and centromeres. This type of heterochromatin is mainly defined epigenetically by specific histone marks such as the di or tri-methylation of lysine 9 of histone H3 (H3K9me3) and the binding of the Heterochromatin protein 1a (HP1a)5,6. On the other hand, facultative heterochromatin localizes through the chromosome&#39;s arms and consists mainly of developmentally silenced genes7,8. Immunostaining of heterochromatin blocks in metaphase cells, or the observation of heterochromatin aggregates in interphase cells, has unveiled much light in the understanding of the formation and function of heterochromatic regions9.
The use of Drosophila as a model system has allowed the development of essential tools to study heterochromatin without the use of electron microscopy10. Since the description of position effect variegation and the discovery of heterochromatin-associated proteins, such as HP1a, and histone post-translational modifications, many groups have developed several immunohistochemical techniques that allow visualization of these heterochromatic regions10,11.
These techniques are based on the use of specific antibodies that recognize heterochromatin-associated proteins or histone marks. For every cell type and antibody, the fixation and permeabilization conditions must be determined empirically. Also, conditions may vary if additional mechanical processes such as squashing techniques are used. In this protocol, we describe the use of Drosophila salivary glands to study heterochromatic foci. Salivary glands have polytenized cells that contain more than 1,000 copies of the genome, thus providing an amplified view of most of the chromatin features, with the exception of satellite DNA and some heterochromatic regions which are under replicated. Nevertheless, heterochromatin regions are easily visualized in polytene chromosome preparations, but the squashing techniques may sometimes disrupt characteristic chromatin-bound complexes or the chromatin architecture. Therefore, immunolocalization of proteins in whole salivary gland tissue can surpass these undesired effects. We have used this protocol to detect several chromatin bound proteins, and we have demonstrated that this protocol combined with mutant Drosophila stocks can be used to study heterochromatin disruption12.

<table><tbody><tr><td>1.5 mL microcentrifuge tubes</td><td>Axygen MCT-150-C</td><td>11351904</td><td>brand not critical</td></tr><tr><td>16% formaldehyde</td><td>Thermo Scientific</td><td>28908</td><td></td></tr><tr><td>AF1 Citifluor</td><td>Ted pella</td><td>19470</td><td>25 mL</td></tr><tr><td>BSA, Molecular Biology Grade</td><td>Roche</td><td>10735078001</td><td>brand not critical</td></tr><tr><td>Complete, protease inhibitors Ultra EDTA-free&lt;br/&gt; protease inhibitors</td><td>Merck</td><td>5892953001</td><td></td></tr><tr><td>Coverslip</td><td>Corning</td><td>CLS285022-200EA</td><td>22x22, brand not critical</td></tr><tr><td>DTT</td><td>Sigma</td><td>d9779</td><td>brand not critical</td></tr><tr><td>EDTA</td><td>Sigma</td><td>E5134</td><td>brand not critical</td></tr><tr><td>EGTA</td><td></td><td></td><td>brand not critical</td></tr><tr><td>Glass slide</td><td>Gold seal</td><td>3011</td><td>brand not critical</td></tr><tr><td>H&lt;sub&gt;3&lt;/sub&gt;BO&lt;sub&gt;3&lt;/sub&gt;</td><td>Baker</td><td>0084-01</td><td>brand not critical</td></tr><tr><td>H3K9me3</td><td>Abcam</td><td>8889</td><td></td></tr><tr><td>HP1a</td><td>Hybridoma Bank</td><td>C1A9</td><td>Product Form&amp;nbsp;Concentrate 0.1 mL</td></tr><tr><td>KCl</td><td>Baker</td><td>3040-01</td><td>brand not critical</td></tr><tr><td>Methanol</td><td>Baker</td><td>9070-03</td><td>brand not critical</td></tr><tr><td>NaCl</td><td>Sigma</td><td>71376</td><td>brand not critical</td></tr><tr><td>NaOH</td><td></td><td></td><td>brand not critical</td></tr><tr><td>PIPES</td><td></td><td></td><td>brand not critical</td></tr><tr><td>Rotator</td><td>Thermo Scientific</td><td>13-687-12Q</td><td>&amp;nbsp;Labquake Tube Shaker</td></tr><tr><td>Thermo Mixer C</td><td>Eppendorf</td><td>13527550</td><td>SmartBlock 1.5 mL</td></tr><tr><td>Tris</td><td>Milipore</td><td>648311</td><td>brand not critical</td></tr><tr><td>Triton X-100</td><td>Sigma</td><td>T8787</td><td>100 mL, brand not critical</td></tr><tr><td>&amp;beta;-mercaptoethanol</td><td>Bio-Rad</td><td>1610710</td><td>25 mL, brand not critical</td></tr></tbody></table>

immunofluorescent staining for visualization of heterochromatin associated proteins in drosophila salivary glands

Visualization of heterochromatin aggregates by immunostaining can be challenging. Many mammalian components of chromatin are conserved in Drosophila melanogaster. Therefore, it is an excellent model to study heterochromatin formation and maintenance. Polytenized cells, such as the ones found in salivary glands of third instar D. melanogaster larvae, provide an excellent tool to observe the chromatin amplified nearly a thousand times and have allowed researchers to study changes in the distribution of heterochromatin in the nucleus. Although the observation of heterochromatin components can be carried out directly in polytene chromosome preparations, the localization of some proteins can be altered by the severity of the treatment. Therefore, the direct visualization of heterochromatin in cells complements this type of study. In this protocol, we describe the immunostaining techniques used for this tissue, the use of secondary fluorescent antibodies, and confocal microscopy to observe these heterochromatin aggregates with greater precision and detail.

Representative results of HP1a immunostaining in Drosophila salivary glands are shown in Figure 1. A positive result is to observe one focal point (Figure 1a) (heterochromatic aggregate or condensate). A negative result is no signal or a dispersed signal. Sometimes a double signal can be observed, that is, with a double point (Figure 1c), but it usually occurs in smaller quantities.
Data analysis...

Watch this Scientific Journal Video about Immunofluorescent Staining for Visualization of Heterochromatin Associated Proteins in Drosophila Salivary Glands at JoVE.com

Immunofluorescent Staining for Visualization of Heterochromatin Associated Proteins in Drosophila Salivary Glands

This protocol aims to visualize heterochromatin aggregates in Drosophila polytene cells.

Immunofluorescent Staining for Visualization of Heterochromatin Associated Proteins in Drosophila Salivary Glands

Visualization of heterochromatin aggregates by immunostaining can be challenging. Many mammalian components of chromatin are conserved in Drosophila ...

immunofluorescent-staining-for-visualization-heterochromatin

Research

JoVE Journal

Biology

3.6K Views.  Universidad Nacional Autónoma de México. This protocol aims to visualize heterochromatin aggregates in Drosophila polytene cells.

Video: Immunofluorescent Staining for Visualization of Heterochromatin Associated Proteins in Drosophila Salivary Glands

Upright Imaging of Drosophila Egg Chambers

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in architecture, is well-characterized, and is amenable to genetic analysis. Each egg chamber consists of germ-line cells surrounded by a single epithelial layer of somatic follicle cells. Subsets of follicle cells undergo differentiation during specific stages to become several different cell types. Standard techniques primarily allow for a lateral view of egg chambers, and therefore a limited view of follicle cell organization and identity. The upright imaging protocol describes a mounting technique that enables a novel, vertical view of egg chambers with a standard confocal microscope. Samples are first mounted between two layers of glycerin jelly in a lateral (horizontal) position on a glass microscope slide. The jelly with encased egg chambers is then cut into blocks, transferred to a coverslip, and flipped to position egg chambers upright. Mounted egg chambers can be imaged on either an upright or an inverted confocal microscope. This technique enables the study of follicle cell specification, organization, molecular markers, and egg development with new detail and from a new perspective.

Upright Imaging of Drosophila Egg Chambers

The upright imaging method described in this protocol allows for the detailed visualization of the poles of a developing Drosophila melanogaster egg. This end-on view provides a new perspective into the arrangements and morphologies of multiple cell types in the follicular epithelium.

Drosophila melanogaster oogenesis provides an ideal context for studying varied developmental processes since the ovary is relatively simple in ...

Developmental Biology

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize mammalian adaptive immunity. Drosophila and mammalian hematopoiesis occur in spatially and temporally distinct phases to produce several blood cell lineages. Both systems maintain reservoirs of blood cell progenitors with which to expand or replace mature lineages. The hematopoietic system allows Drosophila and mammals to respond to and to adapt to immune challenges. Importantly, the transcriptional regulators and signaling pathways that control the generation, maintenance, and function of the hematopoietic system are conserved from flies to mammals. These similarities allow Drosophila to be used to genetically model hematopoietic development and disease.
Here we detail assays to examine the hematopoietic system of Drosophila larvae. In particular, we outline methods to measure blood cell numbers and concentration, visualize a specific mature lineage in vivo, and perform immunohistochemistry on blood cells in circulation and in the hematopoietic organ. These assays can reveal changes in gene expression and cellular processes including signaling, survival, proliferation, and differentiation and can be used to investigate a variety of questions concerning hematopoiesis. Combined with the genetic tools available in Drosophila, these assays can be used to evaluate the hematopoietic system upon defined genetic alterations. While not specifically outlined here, these assays can also be used to examine the effect of environmental alterations, such as infection or diet, on the hematopoietic system.

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Drosophila and mammalian hematopoietic systems share many common features, making Drosophila an attractive genetic model to study hematopoiesis. Here we demonstrate dissection and mounting of the major larval hematopoietic organ for immunohistochemistry. We also describe methods to assay various larval hematopoietic compartments including circulating hemocytes and sessile crystal cells.

Many parallels exist between the Drosophila and mammalian hematopoietic systems, even though Drosophila lack the lymphoid lineage that characterize ...

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

To understand how differences in complex cell shapes are achieved, it is important to accurately follow microtubule organization. The Drosophila larval body wall contains several cell types that are models to study cell and tissue morphogenesis. For example tracheae are used to examine tube morphogenesis1, and the dendritic arborization (DA) sensory neurons of the Drosophila larva have become a primary system for the elucidation of general and neuron-class-specific mechanisms of dendritic differentiation2-5 and degeneration6.
The shape of dendrite branches can vary significantly between neuron classes, and even among different branches of a single neuron7,8. Genetic studies in DA neurons suggest that differential cytoskeletal organization can underlie morphological differences in dendritic branch shape4,9-11. We provide a robust immunological labeling method to assay in vivo microtubule organization in DA sensory neuron dendrite arbor (Figures 1, 2, Movie 1). This protocol illustrates the dissection and immunostaining of first instar larva, a stage when active sensory neuron dendrite outgrowth and branching organization is occurring 12,13.
In addition to staining sensory neurons, this method achieves robust labeling of microtubule organization in muscles (Movies 2, 3), trachea (Figure 3, Movie 3), and other body wall tissues. It is valuable for investigators wishing to analyze microtubule organization in situ in the body wall when investigating mechanisms that control tissue and cell shape.

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

To understand how complex cell shapes, such as neuronal dendrites, are achieved during development, it is important to be able to accurately assay microtubule organization. Here we describe a robust immunohistological labeling method to examine microtubule organization of dendritic arborization neuron sensory dendrites, trachea, muscle, and other Drosophila larva body wall tissues.

To understand how differences in complex cell shapes are achieved, it is important to accurately follow microtubule organization. The Drosophila larval ...

Neuroscience

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Visualizing developing organ formation as well as progession and treatment of disease often heavily relies on the ability to optically interrogate molecular and functional changes in intact living organisms. Most existing optical imaging methods are inadequate for imaging at dimensions that lie between the penetration limits of modern optical microscopy (0.5-1mm) and the diffusion-imposed limits of optical macroscopy (&gt;1cm) [1]. Thus, many important model organisms, e.g. insects, animal embryos or small animal extremities, remain inaccessible for in-vivo optical imaging. 
Although there is increasing interest towards the development of nanometer-resolution optical imaging methods, there have not been many successful efforts in improving the imaging penetration depth. The ability to perform in-vivo imaging beyond microscopy limits is in fact met with the difficulties associated with photon scattering present in tissues. Recent efforts to image entire embryos for example [2,3] require special chemical treatment of the specimen, to clear them from scattering, a procedure that makes them suitable only for post-mortem imaging. These methods however evidence the need for imaging larger specimens than the ones usually allowed by two-photon or confocal microscopy, especially in developmental biology and in drug discovery.
We have developed a new optical imaging technique named Mesoscopic Fluorescence Tomography [4], which appropriate for non-invasive in-vivo imaging at dimensions of 1mm-5mm. The method exchanges resolution for penetration depth, but offers unprecedented tomographic imaging performance and it has been developed to add time as a new dimension in developmental biology observations (and possibly other areas of biological research) by imparting the ability to image the evolution of fluorescence-tagged responses over time. As such it can accelerate studies of morphological or functional dependencies on gene mutations or external stimuli, and can importantly, capture the complete picture of development or tissue function by allowing longitudinal time-lapse visualization of the same, developing organism.
The technique utilizes a modified laboratory microscope and multi-projection illumination to collect data at 360-degree projections. It applies the Fermi simplification to Fokker-Plank solution of the photon transport equation, combined with geometrical optics principles in order to build a realistic inversion scheme suitable for mesoscopic range. This allows in-vivo whole-body visualization of non-transparent three-dimensional structures in samples up to several millimeters in size.
We have demonstrated the in-vivo performance of the technique by imaging three-dimensional structures of developing Drosophila tissues in-vivo and by following the morphogenesis of the wings in the opaque Drosophila pupae in real time over six consecutive hours.

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Mesoscopic fluorescence tomography operates beyond the penetration limits of tissue-sectioning fluorescence microscopy. The technique is based on multi-projection illumination and a photon transport description. We demonstrate in-vivo whole-body 3D visualization of the morphogenesis of GFP-expressing wing imaginal discs in Drosophila melanogaster.

Visualizing  developing organ formation as well as progession and treatment of disease often  heavily relies on the ability to optically interrogate ...

Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling

Drosophila has long been a favorite model system for studying the relationship between chromatin structure and gene regulation due to the cytological advantages provided by the giant salivary gland polytene chromosomes of third instar larvae.  In this tissue the chromosomes undergo many rounds of replication in the absence of cell division giving rise to approximately 1000 copies.  The DNA remains aligned after each replicative cycle resulting in greatly enlarged chromosomes that provide a unique opportunity to correlate chromatin morphology with the localization of specific proteins.  Consequently, there has been a high level of interest in defining the epigenetic modifications present at different genes and at different stages of the transcription process. An important tool for such studies is the labeling of polytene chromosomes with antibodies to the enzyme, transcription factor, or histone modification of interest. This video protocol illustrates the squash technique used in the Johansen laboratory to prepare Drosophila polytene chromosomes for antibody labeling.

This video protocol illustrates the squash technique used in the Johansen laboratory to prepare Drosophila polytene chromosomes for antibody labeling.

Drosophila has long been a favorite model system for studying the relationship between chromatin structure and gene regulation due to the cytological ...

Imaging Centrosomes in Fly Testes

Centrosomes are conserved microtubule-based organelles whose structure and function change dramatically throughout the cell cycle and cell differentiation. Centrosomes are essential to determine the cell division axis during mitosis and to nucleate cilia during interphase. The identity of the proteins that mediate these dynamic changes remains only partially known, and the function of many of the proteins that have been implicated in these processes is still rudimentary. Recent work has shown that Drosophila spermatogenesis provides a powerful system to identify new proteins critical for centrosome function and formation as well as to gain insight into the particular function of known players in centrosome-related processes. Drosophila is an established genetic model organism where mutants in centrosomal genes can be readily obtained and easily analyzed. Furthermore, recent advances in the sensitivity and resolution of light microscopy and the development of robust genetically tagged centrosomal markers have transformed the ability to use Drosophila testes as a simple and accessible model system to study centrosomes. This paper describes the use of genetically-tagged centrosomal markers to perform genetic screens for new centrosomal mutants and to gain insight into the specific function of newly identified genes.

Imaging of centrosomal proteins during Drosophila spermatogenesis is a powerful method to identify new proteins critical for centrosome biology as well as to elucidate the particular function of known players in this process.

Centrosomes are conserved microtubule-based organelles whose structure and function change dramatically throughout the cell cycle and cell ...

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.

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;

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

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Studies performed in Drosophila melanogaster embryos and larvae provide crucial insight into developmental processes such as cell fate specification and organogenesis. Immunostaining allows for the visualization of developing tissues and organs. However, a protective cuticle that forms at the end of embryogenesis prevents permeation of antibodies into late-stage embryos and larvae. While dissection prior to immunostaining is regularly used to analyze Drosophila larval tissues, it proves inefficient for some analyses because small tissues may be difficult to locate and isolate. Sonication provides an alternative to dissection in larval Drosophila immunostaining protocols. It allows for quick, simultaneous processing of large numbers of late-stage embryos and larvae and maintains in situ morphology. After fixation in formaldehyde, a sample is sonicated. Sample is then subjected to immunostaining with antigen-specific primary antibodies and fluorescently labeled secondary antibodies to visualize target cell types and specific proteins via fluorescence microscopy. During the process of sonication, proper placement of a sonicating probe above the sample, as well as the duration and intensity of sonication, is critical. Additonal minor modifications to standard immunostaining protocols may be required for high quality stains. For antibodies with low signal to noise ratio, longer incubation times are typically necessary. As a proof of concept for this sonication-facilitated protocol, we show immunostains of three tissue types (testes, ovaries, and neural tissues) at a range of developmental stages.

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Immunostaining is an effective technique for visualizing specific cell types and proteins within tissues. By utilizing sonication, the protocol described here alleviates the need to dissect Drosophila melanogaster tissues from late-stage embryos and larvae before immunostaining. We provide an efficient methodology for the immunostaining of formaldehyde-fixed whole mount larvae.

Studies performed in Drosophila melanogaster embryos and larvae provide crucial insight into developmental processes such as cell fate specification and ...

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

Drosophila testes are a powerful model system for studying biological processes including stem cell biology, nuclear architecture, meiosis and sperm development. However, immunolabeling of the whole Drosophila testis is often associated with significant non-uniformity of staining due to antibody penetration. Squashed preparations only partially overcome the problem since it decreases the 3D quality of the analyses. Herein, we describe a whole-mount protocol using NP40 and heptane during fixation together with immunolabeling in liquid media. It preserves the volume suitable for confocal microscopy together with reproducible and reliable labeling. We show different examples of 3D reconstitution of spermatocyte nuclei from confocal sections. The intra- and inter-testes reproducibility allows 3D quantification and comparison of fluorescence between single cells from different genotypes. We used different components of the intranuclear MINT structure (Mad1-containing Intra Nuclear Territory) as well as two components associated with the nuclear pore complex to illustrate this protocol and its applications on the largest cells of the testis, the S4-S5 spermatocytes.

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

We present here an improved protocol for the whole-mount preparation and immunostaining of Drosophila testes, suitable for confocal microscopy and also allowing reproducible and reliable labeling. We illustrate this protocol by 3D representation and quantification of colocalization experiments on the large S4-S5 spermatocytes.

Drosophila testes are a powerful model system for studying biological processes including stem cell biology, nuclear architecture, meiosis and sperm ...

Drosophila Larval IHC

Immunohistochemistry (IHC) is a technique used to visualize the presence and location of proteins within tissues. Drosophila larvae are particularly amenable to IHC because of the ease with which they can be processed for staining. Additionally, the larvae are transparent, meaning that some tissues can be visualized without the need for dissection.
In IHC, proteins are ultimately detected with antibodies that specifically bind to &ldquo;epitopes&rdquo; within the protein of interest. In order to preserve these epitopes, tissues must be fixed prior to staining. Furthermore, in order for antibodies to penetrate membranes, cells must be permeabilized with detergents. This video article offers a detailed view of the reagents, tools, and procedures necessary for the staining of dissected larval tissues, including fixation, blocking, and staining steps. Also featured is a demonstration of tissue mounting techniques for fluorescence microscopy. Finally, examples of the broad range of applications for these techniques (and some variations upon them) are provided.

Drosophila melanogaster Larval Immunohistochemistry IHC

Immunohistochemistry (IHC) is a technique used to visualize the presence and location of proteins within tissues. Drosophila larvae are particularly ...

Immunofluorescent Staining for Visualization of Heterochromatin Associated Proteins in Drosophila Salivary Glands

Summary

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Chapters in this video

Upright Imaging of Drosophila Egg Chambers

Methods to Examine the Lymph Gland and Hemocytes in Drosophila Larvae

Immunohistological Labeling of Microtubules in Sensory Neuron Dendrites, Tracheae, and Muscles in the Drosophila Larva Body Wall

Mesoscopic Fluorescence Tomography for In-vivo Imaging of Developing Drosophila

Preparation of Drosophila Polytene Chromosome Squashes for Antibody Labeling

Imaging Centrosomes in Fly Testes

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

Sonication-facilitated Immunofluorescence Staining of Late-stage Embryonic and Larval Drosophila Tissues In Situ

Immunostaining of Whole-Mount Drosophila Testes for 3D Confocal Analysis of Large Spermatocytes

Drosophila Larval IHC