We thank Prof. Eichele for allowing us to use the cryo-microtome. Work was funded by Max-Planck-Gesselschaft.

1. Preparation
<ol>
<li>Freshly prepare Carnoy's fixative by combining absolute ethanol, chloroform and glacial acetic acid in proportion 6:3:1 respectively.3 This, as well as all solutions for submerging collars should be kept in glass staining jars (see reagents section for recommendation).</li>
<li>Prepare aluminum foil pockets the correct size for the collars.</li>
<li>Prepare the following solutions in staining jars: 2 X 40% ethanol, 70% ethanol, 2 X 100% ethanol, methylbenzoate (MB), 50/50 v/v MB and paraffin, 2 X paraffin. Place the MB and paraffin containers in an incubator set to 60-65&deg;C.</li>
<li>Warm paraffin to pour into foil pockets to 60-65&deg;C.</li>
</ol>
2. Fixing Flies in Collars
<ol>
<li>Attach the collar 4 under the binocular using tape in a vertical position where you can see the entry point for the fly.</li>
<li>Anesthetize flies using carbon dioxide or via hypothermia using an ice block. Be careful not to freeze the flies.</li>
<li>Using forceps, pick up individual flies by grabbing their wings and place into the collar oriented properly (head and thorax on top of the blades and abdomen below the blades). 10-20 flies should easily fit into the collar.</li>
</ol>
Note: If you are analyzing several genotypes, do not forget to make a note of the collar number and corresponding genotype.
3. Paraffin Sections of Drosophila Thoraxes
<ol>
<li>Relocate the collar to the Carnoy's solution and fix the tissue at 4&deg;C over night.</li>
<li>After fixation, dehydrate the sample using increasing concentrations of ethanol. For 10 minutes each submerge the collar into 40 % (2 times), 70 % and 100 % (2 times) ethanol at room temperature. Next incubate collar in MB and MB + paraffin solution (1:1) for 30 min in each and then infiltrate the collar in two changes of paraffin for 60 min each at 60-65&deg;C. Rapidly relocate the collar into the foil pocket and fill with melted (60-65&deg;C) paraffin. Place it at room temperature and allow the paraffin to become hard (it is best to leave overnight). Note that the collar can be placed in the foil pocket in different orientations, depending on the orientation of the sections you need (longitudinal or transverse).</li>
<li>Unwrap dry paraffin blocks with collars from the foil and gently separate the collar from the paraffin block. With the help of a sharp blade or scalpel carefully cut out the extra-paraffin from around the fly tissue. </li>
<li>Cut the paraffin block with 7-10 &mu;m section steps on a rotation microtome and allow the cut tissue to float flat in a 37&deg;C water bath. Place the sectioned tissue on polar slides and allow drying over night. These slides can be used for staining with hematoxyline and eosin (Figure 1A-D), toluidine blue, aniline blue or other dies to visualize tissue structures, as well as for antibody staining (Figure 1E-E``).</li>
</ol>
4. Cryosections of Drosophila Thoraxes
<ol>
<li>As with paraffin sections, prepare the flies in a collar and fashion an aluminum foil pocket. A freezing cooler will be needed to prepare the samples. Make sure that the cooler is around -60&deg;C, use a little ethanol and dry ice to allow for reaching this temperature. An hour before starting the experiment put the bottle of cryo-embedding medium (Tissue-Tek O.C.T. compound) upside down in a 4&deg;C fridge in order to cool it down and to minimize the formation of air bubbles.</li>
<li>Relocate the collar with flies to the foil pocket that has been pre-cooled for several minutes inside the freezing cooler and rapidly fill with the cryo-embedding compound. Let the sample freeze for 3-10 min. Carefully unwrap the formed block inside the cooler, gently separate the collar from the embedding block and put it at -20&deg;C for at least one day.</li>
<li>Cut the frozen muscles on a cryo-microtome between -15 and -18&deg;C with a section thickness of 10-15 &mu;m. Place on polarized slides and keep at -20&deg;C until ready to do further processing. We suggest fixing the tissue in 4 % formaldehyde PBS solution for 10 min at room temperature prior to antibody staining.</li>
</ol>
5. Lipids Detection in Drosophila Muscles
Lipid droplets can be detected with oil red O stain on cryosections using a protocol adopted from Sieber and Thummel 5.
<ol>
<li>After fixation, wash slides with water twice for 5 min, equilibrated in propylene glycol for 10 min and incubate for 3 h in oil red O stain at room temperature. Then wash samples 2 times for 5 min in propylene glycol and 30 min in PBS. Mount in 30 % glycerol.</li>
</ol>

6. Representative Results:
<img src="/files/ftp_upload/2438/2438fig1.jpg" alt="Figure 1" /> 
Figure 1. Parrafin-embedded sections 
Hematoxyline and eosine stained transverse (A-B) and longitudinal (C-D) sections of indirect flight muscles. A and C shows normal structured muscles. Abnormal by size and morphology muscles are represented on B and D respectively (black arrows). Transverse section of Drosophila thorax stained with anti-LamC, nuclear envelope marker and DAPI, nuclear stain (E-F). Enlarged view of normal (red arrow) and deteriorated (yellow arrow) muscles (F). G represents section of Drosophila intestinal tract stained with LamC and DAPI.
<img src="/files/ftp_upload/2438/2438fig2.jpg" alt="Figure 2" /> 
Figure 2. Frozen sections 
A. Transverse frozen sections of Drosophila thorax stained with oil red O, lipid droplets label. 
B.	Longitudinal frozen sections of indirect flight muscles stained with anti-Dg, muscle sarcolemma marker and DAPI.

<ol>
	<li>Miller, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+internal+anatomy+and+histology+of+the+imago+of+Drosophila+melanogaster.">The internal anatomy and histology of the imago of Drosophila melanogaster.</a> , (1950).</li><li>Shcherbata, H. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dissecting+muscle+and+neuronal+disorders+in+a+Drosophila+model+of+muscular+dystrophy.">Dissecting muscle and neuronal disorders in a Drosophila model of muscular dystrophy.</a> The EMBO journal. 26, 481-481 (2007).</li><li>Kucherenko, M. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+modifier+screens+reveal+new+components+that+interact+with+the+Drosophila+dystroglycan-dystrophin+complex.">Genetic modifier screens reveal new components that interact with the Drosophila dystroglycan-dystrophin complex.</a> PloS one. 3, e2418-e2418 (2008).</li><li>Puchtler, H., Waldrop, F. S., Conner, H. M., Terry, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Carnoy+fixation:+practical+and+theoretical+considerations.">Carnoy fixation: practical and theoretical considerations.</a> Histochemie. 16, 361-36 (1968).</li><li>Ashburner, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Drosophila - A Laboratory Manual. , (1989).</li><li>Sieber, M. H., Thummel, C. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+DHR96+nuclear+receptor+controls+triacylglycerol+homeostasis+in+Drosophila.">The DHR96 nuclear receptor controls triacylglycerol homeostasis in Drosophila.</a> Cell metabolism. 10, 481-481 (2009).</li></ol>

No conflicts of interest declared.

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<html xmlns="http://www.w3.org/1999/xhtml" xml:lang="en" lang="en">	
		
		<div>
		<table border="1">
		<thead>
		<tr>
		<th>Material Name</th>
		<th>Type</th>
		<th>Company</th>
		<th>Catalogue Number</th>
		<th>Comment</th>
		</tr>
		</thead>
		<tbody>
		
<tr>
<td>Stainless steel collars</td>
<td>&nbsp;</td>
<td>self-made</td>
<td>&nbsp;</td>
<td>Specially constructed</td>
</tr>
<tr>
<td>Forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools GmbH</td>
<td>11295-10</td>
<td>&nbsp;</td>
<tr>
<td>Aluminum foil</td>
<td>&nbsp;</td>
<td>Any</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Blade or scalpel</td>
<td>&nbsp;</td>
<td>Any</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
<tr>
<td>Wheaton macro staining jar</td>
<td>&nbsp;</td>
<td>Wheaton Science Products </td>
<td>900200</td>
<td>&nbsp;</td>
<tr>
<td>Microtome</td>
<td>&nbsp;</td>
<td>Carl Zeiss</td>
<td>&nbsp;</td>
<td>Model: Hyrax M25</td>
</tr>
<tr>
<td>Cryo-microtome</td>
<td>&nbsp;</td>
<td>Leica </td>
<td>&nbsp;</td>
<td>Model CM3050S</td>
</tr>
<tr>
<td>60-65&deg;C Incubator</td>
<td>&nbsp;</td>
<td>Any</td>
<td>&nbsp;</td>
<td>Large enough to hold at least 4 staining jars</td>
</tr>
<tr>
<td>Freezing cooler with metal block</td>
<td>&nbsp;</td>
<td>Any</td>
<td>&nbsp;</td>
<td>Store at -80&deg;C</td>
</tr>
<tr>
<td>Super-frost slides</td>
<td>&nbsp;</td>
<td>Thermo Fisher Scientific</td>
<td>9161155</td>
<td>&nbsp;</td>
<tr>
<td>Cover slips</td>
<td>&nbsp;</td>
<td>Any </td>
<td>&nbsp;</td>
<td>Recommend 24 X 40 mm</td>
</tr>
<tr>
<td>Chloroform</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>288306</td>
<td>Analytical grade </td>
</tr>
<tr>
<td>Glacial acetic acid</td>
<td>&nbsp;</td>
<td>Merck</td>
<td>100063</td>
<td>Analytical grade</td>
</tr>
<tr>
<td>Ethanol</td>
<td>&nbsp;</td>
<td>Merck</td>
<td>100983</td>
<td>Analytical grade</td>
</tr>
<tr>
<td>Methylbenzoate</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>M29908-500G</td>
<td>Analytical grade</td>
</tr>
<tr>
<td>Paraplast plus</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>76258</td>
<td>Paraffin</td>
</tr>
<tr>
<td>Tissue-Teck O.C.T. compound</td>
<td>&nbsp;</td>
<td>Sakura</td>
<td>4583</td>
<td>&nbsp;</td>
<tr>
<td>16% formaldehyde, methanol free</td>
<td>&nbsp;</td>
<td>Polysciences</td>
<td>18814</td>
<td>&nbsp;</td>
<tr>
<td>Glycerol</td>
<td>&nbsp;</td>
<td>Sigma-Aldrich</td>
<td>G5150-1L</td>
<td>&nbsp;</td>		</tbody>
		</table>
		</div>

paraffin-embedded and frozen sections of drosophila adult muscles

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of muscle specification, formation and function in model systems has provided valuable insight into muscle physiology. Therefore, identifying and characterizing molecular mechanisms that underlie muscle damage is critical. The structure of adult Drosophila multi-fiber muscles resemble vertebrate striated muscles 1 and the genetic tractability of Drosophila has made it a great system to analyze dystrophic muscle morphology and characterize the processes affecting muscular function in ageing adult flies 2. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. These preparations allow for the tissue to be stained with classical histological stains and labeled with protein detecting dyes, and specifically cryosections are ideal for immunohistochemical detection of proteins in intact muscles. This allows for analysis of muscle tissue structure, identification of morphological defects, and detection of the expression pattern for muscle/neuron-specific proteins in Drosophila adult muscles. These techniques can also be slightly modified for sectioning of other body parts.

Watch this Scientific Journal Video about Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles at JoVE.com

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

Identification of mechanisms underlying muscle damage is crucial. Here we present the histological technique for preparing paraffin-embedded and frozen sections of Drosophila thoracic muscles. This allows analysis of muscle morphology and localization of protein and other muscle cell components.

Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

The molecular characterization of muscular dystrophies and myopathies in humans has revealed the complexity of muscle disease and genetic analysis of ...

paraffin-embedded-and-frozen-sections-of-drosophila-adult-muscles

Research

JoVE Journal

Biology

36.4K ビュー.  Max Planck Institute for Biophysical Chemistry. 筋肉の損傷のメカニズムの同定は非常に重要です。ここでは、ショウジョウバエの胸部筋肉のパラフィン包埋して凍結切片を調製するための組織学的手法を提示する。これは筋肉の形態やタンパク質と他の筋肉の細胞成分の局在の分析を行うことができます。

ビデオ: Paraffin-Embedded and Frozen Sections of Drosophila Adult Muscles

Whole Cell Recordings from Brain of Adult Drosophila

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by describing the dissecting solution and capture of the adult females used in our studies. The procedure for removing the whole brain intact, including both optic lobes, is illustrated. Dissection of the overlying trachea is also shown.  The isolated brain is not only small but needs special care in handling at this stage to prevent damage to the neurons, many of which are close to the outer surface of the tissue. We show how a special holder we developed is used to stabilize the brain in the recording chamber. A standard electrophysiology set up is used for recording from single neurons or pairs of neurons. A fluorescent image, viewed through the recording microscope, from a GAL4 line driving GFP expression (GH146) illustrates how projection neurons (PNs) are identified in the live brain. A high power Nomarski image shows a view of a single neuron that is being targeted for whole cell recording. When the brain is successfully removed without damage, the majority of the neurons are spontaneously active, firing action potentials and/or exhibiting spontaneous synaptic input. This in situ preparation, in which whole cell recording of identified neurons in the whole brain can be combined with genetic and pharmacological manipulations, is a useful model for exploring cellular physiology and plasticity in the adult CNS.

This video demonstrates the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons using standard whole cell technology. It includes images of GFP labeled cells and neurons viewed during recording.

In this video, we demonstrate the procedure for isolating whole brains from adult Drosophila in preparation for recording from single neurons. We begin by ...

High-Resolution Video Tracking of Locomotion in Adult Drosophila Melanogaster

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field. Studying locomotor behavior in Drosophila melanogaster relies on the ability to be able to quantify changes in motion during or in response to a given task. For this reason, a high-resolution video tracking system, such as the one we describe in this paper, is a valuable tool for measuring locomotion in real-time. Our protocol involves the use of an initial air pulse to break the flies momentum, followed by a thirty second filming period in a square chamber. A tracking program is then used to calculate the instantaneous speed of each fly within the chamber in 10 msec increments. Analysis software then compiles this data, and outputs a variety of parameters such as average speed, max speed, time spent in motion, acceleration, etc. This protocol will discuss proper feeding and management of flies for behavioral tasks, handling flies without anesthetization or immobilization, setting up a controlled environment, and running the assay from start to finish.

The study of complex locomotor behavior in Drosophila melanogaster is dependent upon the ability to quantify changes in a given fly's movement. This article demonstrates how to do this using a high-resolution tracking system.

Flies provide an important model for studying complex behavior due to the plethora of genetic tools available to researchers in this field.  Studying ...

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 ...

Dissection of Oenocytes from Adult Drosophila melanogaster

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

Dissection of Oenocytes from Adult Drosophila melanogaster

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

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

DNA Extraction from Paraffin Embedded Material for Genetic and Epigenetic Analyses

Disease development and progression are characterized by frequent genetic and epigenetic aberrations including chromosomal rearrangements, copy number gains and losses and DNA methylation. Advances in high-throughput, genome-wide profiling technologies, such as microarrays, have significantly improved our ability to identify and detect these specific alterations. However as technology continues to improve, a limiting factor remains sample quality and availability. Furthermore, follow-up clinical information and disease outcome are often collected years after the initial specimen collection. Specimens, typically formalin-fixed and paraffin embedded (FFPE), are stored in hospital archives for years to decades. DNA can be efficiently and effectively recovered from paraffin-embedded specimens if the appropriate method of extraction is applied. High quality DNA extracted from properly preserved and stored specimens can support quantitative assays for comparisons of normal and diseased tissues and generation of genetic and epigenetic signatures 1. To extract DNA from paraffin-embedded samples, tissue cores or microdissected tissue are subjected to xylene treatment, which dissolves the paraffin from the tissue, and then rehydrated using a series of ethanol washes. Proteins and harmful enzymes such as nucleases are subsequently digested by proteinase K. The addition of lysis buffer, which contains denaturing agents such as sodium dodecyl sulfate (SDS), facilitates digestion 2. Nucleic acids are purified from the tissue lysate using buffer-saturated phenol and high speed centrifugation which generates a biphasic solution. DNA and RNA remain in the upper aqueous phase, while proteins, lipids and polysaccharides are sequestered in the inter- and organic-phases respectively. Retention of the aqueous phase and repeated phenol extractions generates a clean sample. Following phenol extractions, RNase A is added to eliminate contaminating RNA. Additional phenol extractions following incubation with RNase A are used to remove any remaining enzyme. The addition of sodium acetate and isopropanol precipitates DNA, and high speed centrifugation is used to pellet the DNA and facilitate isopropanol removal. Excess salts carried over from precipitation can interfere with subsequent enzymatic assays, but can be removed from the DNA by washing with 70% ethanol, followed by centrifugation to re-pellet the DNA 3. DNA is re-suspended in distilled water or the buffer of choice, quantified and stored at -20&deg;C. Purified DNA can subsequently be used in downstream applications which include, but are not limited to, PCR, array comparative genomic hybridization 4 (array CGH), methylated DNA Immunoprecipitation (MeDIP) and sequencing, allowing for an integrative analysis of tissue/tumor samples.

This video demonstrates the protocol for DNA extraction from formalin-fixed paraffin-embedded material. This is a multi-day procedure in which tissue sections are deparaffinized with xylene, rehydrated with ethanol and treated with proteinase K to purify and isolate DNA for subsequent gene-specific or genome-wide analysis.

Disease development and progression are characterized by frequent genetic and epigenetic aberrations including chromosomal rearrangements, copy number ...

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a plethora of mutant strains and technical tricks have been developed to allow sophisticated questions to be asked and answered in a reasonable amount of time. Fundamental insight into the interplay of components of all known major signaling pathways has been obtained in forward and reverse genetic Drosophila studies. The fly eye has proven to be exceptionally well suited for mutational analysis, since, under laboratory conditions, flies can survive without functional eyes. Furthermore, the surface of the insect eye is composed of some 800 individual unit eyes (facets or ommatidia) that form a regular, smooth surface when looked at under a dissecting microscope. Thus, it is easy to see whether a mutation might affect eye development or growth by externally looking for the loss of the smooth surface ('rough eye' phenotype; Fig. 1) or overall eye size, respectively (for examples of screens based on external eye morphology see e.g.1). Subsequent detailed analyses of eye phenotypes require fixation, plastic embedding and thin-sectioning of adult eyes.
The Drosophila eye develops from the so-called eye imaginal disc, a bag of epithelial cells that proliferate and differentiate during larval and pupal stages (for review see e.g. 2). Each ommatidium consists of 20 cells, including eight photoreceptors (PR or R-cells; Fig. 2), four lens-secreting cone cells, pigment cells ('hexagon' around R-cell cluster) and a bristle. The photoreceptors of each ommatidium, most easily identified by their light sensitive organelles, the rhabdomeres, are organized in a trapezoid made up of the six "outer" (R1-6) and two "inner" photoreceptors (R7/8; R8 [Fig. 2] is underneath R7 and thus only seen in sections from deeper areas of the eye). The trapezoid of each facet is precisely aligned with those of its neighbors and the overall anteroposterior and dorsoventral axes of the eye (Fig. 3A). In particular, the ommatidia of the dorsal and ventral (black and red arrows, respectively) halves of the eye are mirror images of each other and correspond to two chiral forms established during planar cell polarity signaling (for review see e.g. 3).
The method to generate semi-thin eye sections (such as those presented in Fig. 3) described here is slightly modified from the one originally described by Tomlinson and Ready4. It allows the morphological analysis of all cells except for the transparent cone cells. In addition, the pigment of R-cells (blue arrowheads in Fig. 2 and 3) can be used as a cell-autonomous marker for the genotype of a R-cell, thus genetic requirements of genes in a subset of R-cells can readily be determined5,6.

Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

A standard approach to prepare adult Drosophila eyes for semi-thin sectioning and light microscopic analysis is presented here. The protocol can be used for gross morphological analysis of eye defects, or with the indicated adjustments can be used to determine genetic requirements of genes in specific cell types of the eye (e.g. clonal analysis of photoreceptors) or for electron microscopic analysis.

Drosophila has long been used as model system to study development, mainly due to the ease with which it is genetically tractable. Over the years, a ...

Proteomic Sample Preparation from Formalin Fixed and Paraffin Embedded Tissue

Preserved clinical material is a unique source for proteomic investigation of human disorders. Here we describe an optimized protocol allowing large scale quantitative analysis of formalin fixed and paraffin embedded (FFPE) tissue. The procedure comprises four distinct steps. The first one is the preparation of sections from the FFPE material and microdissection of cells of interest. In the second step the isolated cells are lysed and processed using 'filter aided sample preparation' (FASP) technique. In this step, proteins are depleted from reagents used for the sample lysis and are digested in two-steps using endoproteinase LysC and trypsin. 
After each digestion, the peptides are collected in separate fractions and their content is determined using a highly sensitive fluorescence measurement. Finally, the peptides are fractionated on 'pipette-tip' microcolumns. The LysC-peptides are separated into 4 fractions whereas the tryptic peptides are separated into 2 fractions. In this way prepared samples allow analysis of proteomes from minute amounts of material to a depth of 10,000 proteins. Thus, the described workflow is a powerful technique for studying diseases in a system-wide-fashion as well as for identification of potential biomarkers and drug targets.

Archival formalin fixed and paraffin embedded (FFPE) clinical samples are valuable material for investigation of diseases. Here we demonstrate a sample preparation workflow allowing in-depth proteomic analysis of microdissected FFPE tissue.

Preserved clinical material is a unique source for proteomic investigation of human disorders. Here we describe an optimized protocol allowing large scale ...

qPCR Is a Sensitive and Rapid Method for Detection of Cytomegaloviral DNA in Formalin-fixed, Paraffin-embedded Biopsy Tissue

It is crucial to identify cytomegalovirus (CMV) infection in the gastrointestinal (GI) tract of immunosuppressed patients, given their greater risk for developing severe infection. Many laboratory methods for the detection of CMV infection have been developed, including serology, viral culture, and molecular methods. Often, these methods reflect systemic involvement with CMV and do not specifically identify local tissue involvement. Therefore, detection of CMV infection in the GI tract is frequently done by traditional histology of biopsy tissue. Hematoxylin and eosin (H&#38;E) staining in conjunction with immunohistochemistry (IHC) have remained the mainstays of examining these biopsies. H&#38;E and IHC sometimes result in atypical (equivocal) staining patterns, making interpretation difficult. It was shown that quantitative polymerase chain reaction (qPCR) for CMV can successfully be performed on formalin-fixed, paraffin-embedded (FFPE) biopsy tissue for very high sensitivity and specificity. The goal of this protocol is to demonstrate how to perform qPCR testing for the detection of CMV in FFPE biopsy tissue in a clinical laboratory setting. This method is likely to be of great benefit for patients in cases of equivocal staining for CMV in GI biopsies.

This protocol describes qPCR detection of cytomegalovirus in formalin-fixed, paraffin-embedded biopsy tissue, which is rapid, sensitive, specific, and useful for interpreting equivocal hematoxylin and eosin or immunohistochemical staining patterns.

It is crucial to identify cytomegalovirus (CMV) infection in the gastrointestinal (GI) tract of immunosuppressed patients, given their greater risk for ...

Indirect Immunofluorescence on Frozen Sections of Mouse Mammary Gland

Indirect immunofluorescence is used to detect and locate proteins of interest in a tissue. The protocol presented here describes a complete and simple method for the immune detection of proteins, the mouse lactating mammary gland being taken as an example. A protocol for the preparation of the tissue samples, especially concerning the dissection of mouse mammary gland, tissue fixation and frozen tissue sectioning, are detailed. A standard protocol to perform indirect immunofluorescence, including an optional antigen retrieval step, is also presented. The observation of the labeled tissue sections as well as image acquisition and post-treatments are also stated. This procedure gives a full overview, from the collection of animal tissue to the cellular localization of a protein. Although this general method can be applied to other tissue samples, it should be adapted to each tissue/primary antibody couple studied.

The indirect immunofluorescence protocol described in this article allows the detection and the localization of proteins in the mouse mammary gland. A complete method is given to prepare mammary gland samples, to perform immunohistochemistry, to image the tissue sections by fluorescence microscopy, and to reconstruct images.

Indirect immunofluorescence is used to detect and locate proteins of interest in a tissue. The protocol presented here describes a complete and simple ...

Employing Digital Droplet PCR to Detect BRAF V600E Mutations in Formalin-fixed Paraffin-embedded Reference Standard Cell Lines

ddPCR is a highly sensitive PCR method that utilizes a water-oil emulsion system. Using a droplet generator, an extracted nucleic acid sample is partitioned into ~20,000 nano-sized, water-in-oil droplets, and PCR amplification occurs in individual droplets. The ddPCR approach is in identifying sequence mutations, copy number alterations, and select structural rearrangements involving targeted genes. Here, we demonstrate the use of ddPCR as a powerful technique for precisely quantitating rare BRAF V600E mutations in FFPE reference standard cell lines, which is helpful in identifying individuals with cancer. In conclusion, ddPCR technique offers the potential to precisely profile the specific rare mutations in different genes in various types of FFPE samples.

The goal of this video is to demonstrate how to perform&#160;automated DNA extraction from formalin-fixed paraffin-embedded (FFPE) reference standard cell lines and digital droplet PCR&#160;(ddPCR) analysis to detect rare mutations in a clinical setting. Detecting mutations in FFPE samples demonstrates the clinical utility of ddPCR in FFPE samples.