I would like to thank Dr. Jennifer Curtiss for the pictures in Fig. 1 and Jeremy Fagan and Dr. Florence Marlow for critically reading the manuscript. Our work is supported by NIH grant 1R01GM088202.

1. Fly head dissection
 <ol>
 <li>Make sure you have all materials at hand (including gelatin coated slides). Prepare glutaraldehyde and Osmium fixatives (see below). Per genotype to embed, aliquot 200&mu;l glutaraldehyde fix solution in 1.5 ml tubes on ice.</li>
 <li>Anesthetize flies on the CO2 pad (we usually dissect seven fly heads to embed six per genotype in case one head is destroyed during the procedure).</li>
 <li>Hold and slightly press the thorax of the fly, dorsal side up, with tweezers and use a sharp scalpel to cut off the head.</li>
 <li>Stabilize the fly head by touching the neck with the tweezers and carefully slice off a small part of one eye. This enhances penetration of the fixative (if you intend to use your eye sections for clonal analysis, cut the eye with no or smaller clones). Avoid touching and damaging the intact eye that will later be sectioned. </li>
 <li>Touch the head with tweezers or the scalpel on the surface of the cut off eye and transfer the head into glutaraldehyde/phosphate fixative on ice (the heads often float on top of the fixative) The dissected heads should not be kept in glutaraldehyde fix for longer than 15 minutes prior to the addition of Osmium. Repeat steps 1.2- 1.5 with the other flies of the same genotype. </li>
 </ol>
2. Fixation and embedding (use gloves for all steps!)
 <ol>
 <li>Spin the flyheads in Eppendorf centrifuge for 1 minute at 10,000 rpm. Continue even if heads don't sink.</li>
 <li>Add 200&mu;l of OsO4 solution and fix for at least 30 minutes and up to 1 hour on ice.</li>
 <li>Using a Pasteur pipette, remove the glutaraldehyde/Osmium solution (make sure to use proper waste disposal procedures!) and replace with 200&mu;l Osmium solution. Incubate on ice for 1-6 hrs. Always ensure the heads are completely covered with liquid and never remove all of the solution as the heads or eyes might collapse!</li>
 <li>Using a Pasteur pipette, discard fixative, add 0.5-1ml 30% ethanol and incubate for at least 5 minutes on ice (make sure to use proper waste disposal procedures since the ethanol now contains Osmium!). </li>
 <li>Repeat the above step with 50%, 70%, (80%), 90% and twice 100% ethanol. Include the 80% ethanol step if using the eyes for electron microscopic analysis. After the 70% wash step, the samples can be removed from the ice.</li>
 <li>Replace the ethanol with an equal volume of propylene oxide (volatile, continue to work in the hood). Incubate for 10 minutes at room temperature.</li>
 <li>Repeat propylene oxide wash. Work in batches if processing many genotypes in parallel to avoid collapsing of the eyes due to evaporation of the propylene oxide.</li>
 <li>Remove most of the propylene oxide and using a plastic transfer pipette, add about 500&mu;l of a 1:1 resin:propylene oxide solution. Equilibrate overnight at room temperature. </li>
 </ol>
3. Embedding and arrangement in molds
 <ol>
 <li>Make sure that all molds are labeled to keep track of your different genotypes, then fill the molds with 100% resin. Do not overfill (no 'high hills' over the surface of the molds when looking straight across the mold surface). Use hard resin for EM analysis. </li>
 <li>Using a transfer pipette, remove the 50% resin from the heads, replace with 100% resin, and incubate for 3-4 hours at room temperature. As heads are infiltrated with resin, they will sink to the bottom of the tube.</li>
 <li>Using a toothpick that has been hit on a hard surface once to create a little hook, transfer one head at the time into a mold. </li>
 <li>Place aluminum foil on the working area of your dissecting scope to protect it against resin spills.</li>
 <li>Using a dissecting needle and looking though your scope, swirl the head slightly in the resin and carefully move it to the bottom of the mold close to the pointed end.</li>
 <li>Carefully align the surface of the intact (!) eye (or your future sectioning plane if you want to have cross-sections) neck down with the front wall of the mold, making sure to keep the tangential surface of the eye roughly half a head diameter away from the mold wall. In rare cases, an eye collapses which can be seen as an empty space between the eye surface and the head cuticle. If you have a collapsed eye, replace it with the 7th spare eye. </li>
 <li>Repeat with all heads. Once done with all alignments, re-check all eyes to make sure they did not move when you removed the needle out of the resin.</li>
 <li>If you have trouble with eyes moving when you remove the dissecting needle, position the eye as desired, quickly retract the needle slightly away from the head and then slowly remove the needle completely.</li>
 <li>Bake the molds overnight at 70&deg;C.</li>
 </ol>
4. Trimming and sectioning 
 <ol>
 <li>Remove hardened blocks from the molds by bending the molds, keeping track of which block corresponds to which genotype (e.g. store in small scintillation vials).</li>
 <li>For trimming, use a chuck appropriate for your microtome that is mounted face up on a block of Plexiglas or aluminum. Mount the block to trim, eye up, in the chuck. </li>
 <li>Under the dissection scope using a Teflon coated razor bade, carefully cut off only the top layer of your block. This will leave a clear surface though which you should see the eye and can thus predict the future plane of sectioning. Avoid cutting your fingers!</li>
 <li>Cut off excess plastic on both sides of the head and on the front (i.e. what used to be the top of the mold when aligning the eyes) until most of the plastic around the head is cut off. It is best to slowly approach the head by cutting off multiple thin layers of plastic.</li>
 <li>Using a clean razor blade, carefully remove thin layers from the top of the eye in the exact plane you want to section later (usually tangentially to the eye, at a slight angle to the original mold surface).</li>
 <li>Continue until you just start to cut away the outside surface of the eye. Make sure you use clean areas of the blade to obtain a shiny, transparent surface, which makes alignment in the microtome easier.</li>
 <li>To save on razor blades, use older blades for the crude trimming around the sides of the block and use a fresh blade for the fine trimming (first cut and 4.6). In the end you will have a three sided pyramid with a slightly tilted, cut-off top that corresponds to your future section plane.</li>
 <li>Mount the block in the microtome. Actual sectioning will vary depending on the type of microtome and will not be described in detail here. We use a Sorvall MT5000 with a histo quality diamond knife to cut 0.5 to 1&mu;m sections. For light microscopic analysis, slight variations in section thickness do not matter. If sectioning clones marked by a White+ transgene, section 1&mu;m sections to ensure to have enough pigment granules adjacent to the rhabdomeres per section (Fig. 2).</li>
 <li>Sections will float on water. Lift first 20-30 sections with a flattened end of a wooden Q-tip from underneath the water surface in a rotating movement and transfer them into a drop of water on a gelatin coated glass slide (kept on a heating plate at around 100&deg;C).</li>
 <li>Repeat with another 20-30 sections. Wait until water has evaporated and sections stick to the slide. Usually, sections of 3 eyes arranged in three columns (three eyes) by twp rows (top/bottom sections) fit well on one slide.</li>
 </ol>
5. Staining and microscopic analysis
 <ol>
 <li>Unless sectioning clones, stain sections. Place slide with sections on a heating block set at 90&deg;C. Dispense Toluidine-blue staining solution from a 30 ml syringe attached to a 0.22&mu;m filter onto the slide and stain for 10 seconds. Immediately rinse extensively with distilled water. Air dry.</li>
 <li>Using a plastic transfer pipette, distribute 3 drops of DPX mounting medium onto the slide and cover with a coverslip. Top with 3 dimes and let the mounting medium harden (e.g. overnight at room-temperature).</li>
 <li>Image with a light microscope. Use a 5x or 10x lens to find a good section and then switch to a 63x oil immersion lens for detailed analysis and photographing. We usually use phase contrast settings, but darkfield optics work as well.</li>
 </ol>
6. Representative results:
Most frequently, eyes are embedded in plastic for a detailed analysis of external rough eye phenotypes detected as part of a genetic screen or in the process of testing genetic interactions with genes known to affect either general eye structure or polarity. Typical results of eye sections are shown in Figure 3. Wild-type ommatidia (Fig. 3A) show the full photoreceptor complement surrounded by a lattice of pigment cells. In contrast, in a strabismus (stbm, a.k.a. van-gogh7) mutant, the ommatidial polarity is lost and, even though the full photoreceptor complement is present, rotation and chirality are randomized (Fig. 3B). Furthermore, since the stbm allele sectioned was in a w- background, the pigment granules are missing in Fig. 3B. Figure 3C shows an example of a clonal analysis. Drosophila Rho kinase (drok) is required for ommatidial rotation as well as for the structural integrity of the eye8. Homozygous drok2 clones can be identified by the absence of pigment in the pigment cells and the rhabdomeres (typically, clones are induced using eyeless-FLP and a P-element with a strongly expressed white marker gene complementing a background w- mutation in the wild-type and heterozygous tissue). It is thus possible to identify mutant areas by their lack of pigment. Due to the cell-autonomy of the pigmentation in the R-cells mentioned earlier, genotypes of individual R-cells can be determined (see arrowheads in Fig. 3C).
<img src="/files/ftp_upload/2959/2959fig1.jpg" alt="Figure 1"/> Figure 1. The Drosophila eye is an excellent model system for biologists since, in contrast to the smooth surface of a wild-type eye (A), the surface of a mutant eye is externally frequently rough (B), indicative of underlying eye phenotypes. In all figures, anterior is to the left and dorsal is up. Images Courtesy of Dr. Jennifer Curtiss, NMSU, Las Cruces, NM, USA. 
<img src="/files/ftp_upload/2959/2959fig2.jpg" alt="Figure 2"/> Figure 2. Light microscopic images of single ommatidia sectioned using the described method. Only seven R-cells are visible at a time per ommatidium, since R7 lies on top of R8. (A, A') On the R7 level, the cell body of R7 is detected between R1 and R6 (yellow arrow in A'). In contrast, at the R8 level (B), the cell body of R8 is detected between R1 and R2 (yellow arrow in B'). Blue arrowheads mark the pigment granules that can be used as a cell-autonomous pigment marker.
<img src="/files/ftp_upload/2959/2959fig3.jpg" alt="Figure 3"/> Figure 3. Tangential sections through wild-type (A), stbm (B) and drok (C) mutant adult Drosophila eyes. Schematics below the sections indicate the polarity of ommatidia (see (A) for arrows). Circles represent ommatidia with defects in the photoreceptor complement. Yellow lines represent the dorsal/ventral line of symmetry (equator). In contrast to the well-oriented ommatidia of wild-type (A), the planar organization is lost in the stbm mutant (B). Note that the section of the stbm mutant shown lacks pigment due to its w- mutant background. (C) Because drok mutations are lethal, they have to be analyzed in clones. Thus, pigmented cells are wild-type or heterozygous (filled blue arrowheads), while R-cells lacking pigment (open blue arrowheads) are homozygous mutant. In addition to rotation defects, drok mutants also show structural defects including missing or excess numbers of R-cells. Note that for clonal analysis, the sections are not stained to avoid obscuring the pigment granules.

<ol>
	<li>Harvey, K. F., Pfleger, C. M., Hariharan, I. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Drosophila+Mst+ortholog,+hippo,+restricts+growth+and+cell+proliferation+and+promotes+apoptosis.">The Drosophila Mst ortholog, hippo, restricts growth and cell proliferation and promotes apoptosis.</a> Cell. 114, 457-457 (2003).</li><li>Secombe, J., Li, L., Carlos, L., Eisenman, R. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Trithorax+group+protein+Lid+is+a+trimethyl+histone+H3K4+demethylase+required+for+dMyc-induced+cell+growth.">The Trithorax group protein Lid is a trimethyl histone H3K4 demethylase required for dMyc-induced cell growth.</a> Genes Dev. 21, 537-537 (2007).</li><li>Cagan, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Principles+of+Drosophila+eye+differentiation.">Principles of Drosophila eye differentiation.</a> Curr Top Dev Biol. 89, 115-115 (2009).</li><li>Jenny, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Planar+cell+polarity+signaling+in+the+Drosophila+eye.">Planar cell polarity signaling in the Drosophila eye.</a> Curr Top Dev Biol. 93, 189-189 (2010).</li><li>Tomlinson, A., Ready, D. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell+fate+in+the+Drosophila+Ommatidium.">Cell fate in the Drosophila Ommatidium.</a> Dev. Biol. 123, 264-264 (1987).</li><li>Tomlinson, A., Ready, D. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neuronal+differentiation+in+the+Drosophila+ommatidium.">Neuronal differentiation in the Drosophila ommatidium.</a> Dev. Biol. 120, 366-366 (1987).</li><li>Jenny, A., Darken, R. S., Wilson, P. A., Mlodzik, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Prickle+and+Strabismus+form+a+functional+complex+to+generate+a+correct+axis+during+planar+cell+polarity+signaling.">Prickle and Strabismus form a functional complex to generate a correct axis during planar cell polarity signaling.</a> Embo J. 22, 4409-4409 (2003).</li><li>Zheng, L., Zhang, J., Carthew, R. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=frizzled+regulates+mirror-symmetric+pattern+formation+in+the+Drosophila+eye.">frizzled regulates mirror-symmetric pattern formation in the Drosophila eye.</a> Development. 121, 3045-3045 (1995).</li><li>Reinke, R., Zipursky, S. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cell-cell+interaction+in+the+Drosophila+Retina:+The+bride+of+sevenless+Gene+is+required+in+photoreceptor+cell+R8+for+R7+cell+development.">Cell-cell interaction in the Drosophila Retina: The bride of sevenless Gene is required in photoreceptor cell R8 for R7 cell development.</a> Cell. 55, 321-321 (1988).</li><li>Wolff, T., Rubin, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=strabismus,+a+novel+gene+that+regulates+tissue+polarity+and+cell+fate+decisions+in+Drosophila.">strabismus, a novel gene that regulates tissue polarity and cell fate decisions in Drosophila.</a> Development. 125, 1149-1149 (1998).</li><li>Winter, C. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drosophila+Rho-associated+kinase+(Drok)+links+Frizzled-mediated+planar+cell+polarity+signaling+to+the+actin+cytoskeleton.">Drosophila Rho-associated kinase (Drok) links Frizzled-mediated planar cell polarity signaling to the actin cytoskeleton.</a> Cell. 105, 81-81 (2001).</li><li>Xu, T., Rubin, G. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+genetic+mosaics+in+developing+and+adult+Drosophila+tissues.">Analysis of genetic mosaics in developing and adult Drosophila tissues.</a> Development. 117, 1223-1223 (1993).</li><li>Stocker, H., Gallant, P., Dahmann, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Methods Mol Biol. 420, 27-27 (2008).</li></ol>

No conflicts of interest declared.

Using Drosophila as model organism, genetic screens led to the identification of many of the founding members of gene families essential for most of the highly conserved signaling pathways in higher eukaryotes including humans. Since, under laboratory conditions, a functional eye is dispensable for survival, the eye is a particularly well suited tissue for the discovery of novel gene functions and the assessment of genetic networks. Ultrastructural analysis of the fly eye using the described method thus led to fundamental discoveries relevant for development and disease. Initially, single cell clonal analysis was performed using X-ray induced clones combined with known closely associated cell-autonomous recessive markers. More recently, the availability of the FLP/FRT system to generate clones has greatly facilitated the phenotypic analysis of lethal mutations in eye sections6,9.
Analysis of eye sections is not limited to tangential sections described here. If desired, the heads can be aligned in any orientation in the molds and transverse sections can be obtained to study deeper layers in the head such as the lamina and medulla. The protocol described here is thus a versatile method for analyses of eye and head structures Drosophila adults.

General equipment:
	<ul>
			<li>For general fly husbandry see 10</li>
			<li>Standard dissection microscope (e.g. Nikon SMZ-645 Stereo Zoom Microscope), preferably with a ring light source to prevent casting a shadow of your hands. </li>
			<li>CO2 Fly pad, covered with 3MM Whatman paper to provide protection against dirt during dissection. </li>
			<li>Gelatin coated slides: dissolve 10g gelatin and 1g CrKSO4x 12 H2O in 1L water on a heating plate (bring almost to a boil, takes approximately 2 hrs). Dip well washed (soap) and rinsed (dH2O) slides in holders into gelatin solution and air-dry covered with foil overnight.</li>
			<li>Heating oven (70&deg;C; old vacuum ovens that were used for nitrocellulose filter drying are well suited for this purpose).</li>
			<li>Microtome capable of ultrathin sectioning (0.5-1 &mu;m thick sections), e.g. those used by electron microscopy facilities. We use a Sorvall MT5000 Ultramicrotome with a Histo quality diamond knife. Glass knives also work.</li>
	</ul>
Fixation solutions:
	<ul>
			<li>Prepare a stock solution of 0.2 M Sodium Phosphate buffer, pH 7.2.</li>
			<li>Prepare 2% glutaraldehyde in 0.1M Sodium Phosphate buffer, pH 7.2 (can be stored at 4&deg;C for 4 weeks). Make at least enough to have 200&mu;l per genotype you want to embed. Keep on ice.</li>
			<li>Osmium solution: 2% OsO4 in 0.1M Sodium Phosphate buffer, pH 7.2. Make only enough to have 200&mu;l per genotype you want to embed, since the solution cannot be stored. Keep on ice.</li>
	</ul>
Glutaraldehyde and OsO4 are highly toxic and should be handled with extreme care in a well-ventilated hood. Use filter tips when handling OsO4 to prevent blackening of your pipette.
Dehydration: Make 30%, 50%, 70%, (80%), 90%, and 100% ethanol solutions. Preferably cool 30%, 50%, 70% ethanol on ice before use.
Staining solution:
1% Toluidine-blue in 1% Borax.
<table border="1">
	<tbody>
		<tr>
			<th>&nbsp;</th>
			<th>Soft</th>
			<th>Hard</th>
		</tr>
		<tr>
			<td>Resin A</td>
			<td>54g</td>
			<td>50g</td>
		</tr>
		<tr>
			<td>Hardener B</td>
			<td>44.5g</td>
			<td>50g</td>
		</tr>
		<tr>
			<td>Accelerator C</td>
			<td>2.5g</td>
			<td>1.75g</td>
		</tr>
		<tr>
			<td>Plasticizer D</td>
			<td>10g</td>
			<td>0.75g</td>
		</tr>
	</tbody>
</table>
Table 1. Resin preparation
To prepare the resin, carefully, but thoroughly mix all ingredients in a plastic beaker using a magnetic stirrer with a large stir bar. Avoid bubbles. After mixing, aliquot in standard scintillation vials or similar containers and freeze at -20&deg;C. Use soft resin for all applications unless your EM facility asks for the harder formulation. Use gloves to handle resin (carcinogenic in its unpolymerized form). To polymerize resin on equipment and waste, bake at 70&deg;C overnight. Waste may then be safely discarded and equipment cleaned and reused. Spilled unpolymerized resin can be cleaned with isopropanol.
<table border="1">
	<tbody>
		<tr>
			<th>Reagent</th>
			<th>Supplier</th>
			<th>Catalog number</th>
		</tr>
		<tr>
			<td>Fly pad	</td>
			<td>e.g. Genesee</td>
			<td>59-119</td>
		</tr>
		<tr>
			<td>Glutaraldehyde</td>
			<td>Sigma</td>
			<td>G7526-10 X 10 ML</td>
		</tr>
		<tr>
			<td>OsO4</td>
			<td>Polysciences,Inc.</td>
			<td>0972A-20</td>
		</tr>
		<tr>
			<td>Propyleneoxide</td>
			<td>Fisher</td>
			<td>04332-1</td>
		</tr>
		<tr>
			<td>Scalpel handles, for #3</td>
			<td>Fisher</td>
			<td>22080046</td>
		</tr>
		<tr>
			<td>Scalpel blades #11	</td>
			<td>Fisher</td>
			<td>08-916-5B</td>
		</tr>
		<tr>
			<td>Transfer pipettes</td>
			<td>Fisher</td>
			<td>13-711-7M</td>
		</tr>
		<tr>
			<td>Durcupan (R) ACM resin</td>
			<td>Sigma (Fluka)</td>
			<td>44610-1EA</td>
		</tr>
		<tr>
			<td>Steel dissecting needle</td>
			<td>Fisher</td>
			<td>S17346</td>
		</tr>
		<tr>
			<td>BEEM flat embedding mold</td>
			<td>Electron Microscopy Sci.</td>
			<td>70904-12</td>
		</tr>
		<tr>
			<td>Teflon coated razor blades</td>
			<td>Electron Microscopy Sci.</td>
			<td>71970</td>
		</tr>
		<tr>
			<td>Q-tip (sterile swabs)</td>
			<td>Fisher</td>
			<td>14-959-81</td>
		</tr>
		<tr>
			<td>Glass slides</td>
			<td>Fisher</td>
			<td>12-550-143</td>
		</tr>
		<tr>
			<td>Cover slips No 1, 22X60 mm</td>
			<td>Fisher</td>
			<td>12-531K</td>
		</tr>
		<tr>
			<td>Gelatin</td>
			<td>Fisher</td>
			<td>ICN96010280</td>
		</tr>
		<tr>
			<td>Cr(III) K SO4 dodecahydrate</td>
			<td>Sigma</td>
			<td>243361</td>
		</tr>
		<tr>
			<td>Diamond Histo knife, 6mm</td>
			<td>Diatome US</td>
			<td>60-His</td>
		</tr>
		<tr>
			<td>Toluidine Blue O</td>
			<td>Fisher	</td>
			<td>BP107-10</td>
		</tr>
		<tr>
			<td>Borax (Na-Tetraborate)</td>
			<td>Fisher</td>
			<td>AC20629-1000</td>
		</tr>
		<tr>
			<td>DPX mounting medium</td>
			<td>Sigma</td>
			<td>44581-100ML</td>
		</tr>
	</tbody>
</table>
Table 2. Materials

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.

Watch this Scientific Journal Video about Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis at JoVE.com

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.

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 ...

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16.1K Views.  Albert Einstein College of Medicine. Стандартный подход к подготовке взрослых Drosophila Глаза для полу-тонкие и легкие срезов микроскопического анализа представлена ​​здесь. Протокол может быть использован для валового морфологического анализа глазом дефектов, или с указанной поправки м...

Video: Preparation of Adult Drosophila Eyes for Thin Sectioning and Microscopic Analysis

MALDI Sample Preparation: the Ultra Thin Layer Method

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS) 1,2. The ultra-thin layer method involves the production of a substrate layer of matrix crystals (alpha-cyano-4-hydroxycinnamic acid) on the sample plate, which serves as a seeding ground for subsequent crystallization of a matrix/analyte mixture. Advantages of the ultra-thin layer method over other sample deposition approaches (e.g. dried droplet) are that it provides (i) greater tolerance to impurities such as salts and detergents, (ii) better resolution, and (iii) higher spatial uniformity. This method is especially useful for the accurate mass determination of proteins. The protocol was initially developed and optimized for the analysis of membrane proteins and used to successfully analyze ion channels, metabolite transporters, and receptors, containing between 2 and 12 transmembrane domains 2. Since the original publication, it has also shown to be equally useful for the analysis of soluble proteins. Indeed, we have used it for a large number of proteins having a wide range of properties, including those with molecular masses as high as 380 kDa 3. It is currently our method of choice for the molecular mass analysis of all proteins. The described procedure consistently produces high-quality spectra, and it is sensitive, robust, and easy to implement.

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption Ionization Mass Spectrometry (MALDI-MS).

This video demonstrates the preparation of an ultra-thin matrix/analyte layer for analyzing peptides and proteins by Matrix-Assisted Laser Desorption ...

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 ...

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 ...

Dissection of Hippocampal Dentate Gyrus from Adult Mouse

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its remarkable neuronal cell plasticity, and its involvement in epilepsy, neurodegenerative diseases, and psychiatric disorders. The hippocampus is composed of distinct regions; the dentate gyrus, which comprises mainly granule neurons, and Ammon's horn, which comprises mainly pyramidal neurons, and the two regions are connected by both anatomic and functional circuits. Many different mRNAs and proteins are selectively expressed in the dentate gyrus, and the dentate gyrus is a site of adult neurogenesis; that is, new neurons are continually generated in the adult dentate gyrus. To investigate mRNA and protein expression specific to the dentate gyrus, laser capture microdissection is often used. This method has some limitations, however, such as the need for special apparatuses and complicated handling procedures. In this video-recorded protocol, we demonstrate a dissection technique for removing the dentate gyrus from adult mouse under a stereomicroscope. Dentate gyrus samples prepared using this technique are suitable for any assay, including transcriptomic, proteomic, and cell biology analyses. We confirmed that the dissected tissue is dentate gyrus by conducting real-time PCR of dentate gyrus-specific genes, tryptophan 2,3-dioxygenase (TDO2) and desmoplakin (Dsp), and Ammon's horn enriched genes, Meis-related gene 1b (Mrg1b) and TYRO3 protein tyrosine kinase 3 (Tyro3). The mRNA expressions of TDO2 and Dsp in the dentate gyrus samples were detected at obviously higher levels, whereas Mrg1b and Tyro3 were lower levels, than those in the Ammon's horn samples. To demonstrate the advantage of this method, we performed DNA microarray analysis using samples of whole hippocampus and dentate gyrus. The mRNA expression of TDO2 and Dsp, which are expressed selectively in the dentate gyrus, in the whole hippocampus of alpha-CaMKII+/- mice, exhibited 0.037 and 0.10-fold changes compared to that of wild-type mice, respectively. In the isolated dentate gyrus, however, these expressions exhibited 0.011 and 0.021-fold changes compared to that of wild-type mice, demonstrating that gene expression changes in dentate gyrus can be detected with greater sensitivity. Taken together, this convenient and accurate dissection technique can be reliably used for studies focused on the dentate gyrus.

A dissection technique for removal of the dentate gyrus from adult mouse under a stereomicroscope was demonstrated in this video-recorded protocol.

The hippocampus is one of the most widely studied areas in the brain because of its important functional role in memory processing and learning, its ...

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 ...

Preparation of Drosophila S2 cells for Light Microscopy

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature and genome sequence database. Cultured Drosophila S2 cells, the product of disassociated 20-24 hour old embryos1, possess all these properties. Consequently, S2 cells are extremely well-suited for the analysis of cellular processes, including the discovery of the genes encoding the molecular components of the process or mechanism of interest. The features of S2 cells that are most responsible for their utility are the ease with which they are maintained, their exquisite sensitivity to double-stranded (ds)RNA-mediated interference (RNAi), and their tractability to fluorescence microscopy as either live or fixed cells.
S2 cells can be grown in a variety of media, including a number of inexpensive, commercially-available, fully-defined, serum-free media2. In addition, they grow optimally and quickly at 21-24&deg;C and can be cultured in a variety of containers. Unlike mammalian cells, S2 cells do not require a regulated atmosphere, but instead do well with normal air and can even be maintained in sealed flasks.
Complementing the ease of RNAi in S2 cells is the ability to readily analyze experimentally-induced phenotypes by phase or fluorescence microscopy of fixed or live cells. S2 cells grow in culture as a single monolayer but do not display contact inhibition. Instead, cells tend to grow in colonies in dense cultures. At low density, S2 cultures grown on glass or tissue culture-treated plastic are round and loosely-attached. However, the cytology of S2 cells can be greatly improved by inducing them to flatten extensively by briefly culturing them on a surface coated with the lectin, concanavalin A (ConA)3. S2 cells can also be stably transfected with fluorescently-tagged markers to label structures or organelles of interest in live or fixed cells. Therefore, the usual scenario for the microscopic analysis of cells is this: first, S2 cells (which can possess transgenes to express tagged markers) are treated by RNAi to eliminate a target protein(s). RNAi treatment time can be adjusted to allow for differences in protein turn-over kinetics and to minimize cell trauma/death if the target protein is important for viability. Next, the treated cells are transferred to a dish containing a coverslip pre-coated with conA to induce cells to spread and tightly adhere to the glass. Finally, cells are imaged with the researcher's choice of microscopy modes. S2 cells are particularly good for studies requiring extended visualization of live cells since these cells stay healthy at room temperature and normal atmosphere.

Preparation of Drosophila S2 cells for Light Microscopy

Drosophila Schneider (S2) cells are an increasingly popular system for the discovery and functional analysis of genes. Our goal is to describe some of the microscopic techniques that make S2 cells such an increasingly important experimental system.

The ideal experimental system would be cheap and easy to maintain, amenable to a variety of techniques, and would be supported by an extensive literature ...

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.

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.

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

Laser Capture Microdissection of Highly Pure Trabecular Meshwork from Mouse Eyes for Gene Expression Analysis

Laser capture microdissection (LCM) has allowed gene expression analysis of single cells and enriched cell populations in tissue sections. LCM is a great tool for the study of the molecular mechanisms underlying cell differentiation and the development and progression of various diseases, including glaucoma. Glaucoma, which comprises a family of progressive optic neuropathies, is the most common cause of irreversible blindness worldwide. Structural changes and damage within the trabecular meshwork (TM) can result in increased intraocular pressure (IOP), which is a major risk factor for developing glaucoma. However, the precise molecular mechanisms involved are still poorly understood. The ability to perform gene expression analysis will be crucial in obtaining further insights into the function of these cells and its role in the regulation of IOP and glaucoma development. To achieve this, a reproducible method for isolating highly enriched TM from frozen sections of mouse eyes and a method for downstream gene expression analysis, such as RT-qPCR and RNA-Seq is needed. The method described herein is developed to isolate highly pure TM from mouse eyes for downstream digital PCR and microarray analysis. In addition, this technique can be easily adapted for the isolation of other highly enriched ocular cells and cell compartments that have been difficult to isolate from mouse eyes. The combination of LCM and RNA analysis can contribute to a more comprehensive understanding of the cellular events underlying glaucoma.

Here, we describe a protocol for a reproducible laser capture microdissection (LCM) for isolating trabecular meshwork (TM) for downstream RNA analysis. The ability to analyze changes in gene expression in the TM will help in understanding the underlying molecular mechanisms of TM-related ocular diseases.

Laser capture microdissection (LCM) has allowed gene expression analysis of single cells and enriched cell populations in tissue sections. LCM is a great ...

Automated Charting of the Visual Space of Housefly Compound Eyes

This paper describes the automatic measurement of the spatial organization of the visual axes of insect compound eyes, which consist of several thousands of visual units called ommatidia. Each ommatidium samples the optical information from a small solid angle, with an approximate Gaussian-distributed sensitivity (half-width on the order of 1&#730;) centered around a visual axis. Together, the ommatidia gather the visual information from a nearly panoramic field of view. The spatial distribution of the visual axes thus determines the eye's spatial resolution. Knowledge of the optical organization of a compound eye and its visual acuity is crucial for quantitative studies of neural processing of the visual information. Here we present an automated procedure for mapping a compound eye's visual axes, using an intrinsic, in vivo optical phenomenon, the pseudopupil, and the pupil mechanism of the photoreceptor cells. We outline the optomechanical setup for scanning insect eyes and use experimental results obtained from a housefly, Musca domestica, to illustrate the steps in the measurement procedure.

The protocol here describes the measurement of the spatial organization of the visual axes of housefly eyes, mapped by an automatic device, using the pseudopupil phenomenon and the pupil mechanism of the photoreceptor cells.

This paper describes the automatic measurement of the spatial organization of the visual axes of insect compound eyes, which consist of several thousands ...