This work was supported by an Ehrman fellowship to H-Y. H., a Jane Coffin Childs Memorial Fund for Medical Research postdoctoral fellowship to R.J.J., NIH Grant F32EY016309 to D.V., a New York University Dean's Dissertation Fellowship to D.J., NIH GrantR01 EY13010 to C.D. and a DFG fellowship to J.R. (RI 2208/1-1). We thank Nina Vogt and Pamela Boodram for comments on the manuscript.

1. Introduction
In this video, we describe methods for retinal dissections and immunohistochemistry at three defined developmental periods: the third instar larval, the midpupal and the adult stage. Although our protocol also works for other pupal stages (for details on earlier stages, see 16), we chose the midpupal stage, as it is optimal for imaging all PRs in one focal plane and their nuclei are easily identifiable, which facilitates the visualization of transcription factor expression patterns. The dissection of late pupal retinas is very similar to the dissection of adult retinas.
First, we demonstrate how to collect larvae and pupae at the desired stages. Then, we explain how to obtain optimal dissections of larval 17-18, midpupal and adult retinas. Second, we demonstrate how to visualize PR development at these stages using immunohistochemistry and confocal microscopy.
2. Staging and Preparation of Specimen
The developmental period of insects is temperature-dependent. To facilitate reproducible staging of larvae and pupae, we recommend raising all fly stocks at 25 &deg;C under a 12 hr/12 hr light/dark cycle.
Third-instar larvae: About 96 hr after egg laying (at 25 &deg;C), late third instar larvae stop feeding and wander on the walls of the food vial, where they eventually pupate. Use a soft brush or a forceps to gently remove a wandering third-instar larva from the wall of the food vial.
50% pupae ('midpupae') are observed about 48 hr after pupation. For exact timing, you can circle just-formed white pupae with a marker on the food vial. Alternatively, move all white pupae to a new vial to obtain a vial of uniform staging (for staging of pupae using morphological criteria, see 19). After 48 hr, carefully remove the midpupae from the vial using a forceps.
Adult flies: Anesthetize young adult flies (2-5 days after eclosion) on a CO2 pad. Remove heads using a forceps and store them in a glass well dish containing cold 1x PBS.
3. Dissection Procedure (About 1-2 hr per Experiment)
After collecting the specimen, place them in a drop of cold PBS on a Sylgard dissection dish. Dissect about 10-15 retinas per genotype.
Please consult the Material Safety Data Sheets as well as your institution's Environmental Health and Safety Office for proper handling of the reagents and equipment (see tables at the end of this protocol).
1. Larval Eye Imaginal Discs
<ol>
	<li>Locate the anterior end of the larva, which contains the mouth hooks. Use a forceps to press the middle part of the larva gently against the base of the Sylgard dish. Use another forceps to grab the mouth hooks and pull gently and slowly away from the body. Discard the main part of the body and use the mouthparts to hold the anterior part in order to remove extraneous tissue from the imaginal discs that remain attached to the brain.</li>
	<li>Transfer brain with imaginal discs by grabbing the mouth hooks with a forceps to 1x PBS in a three-well glass dish. Perform fixation step (step 4.1.).</li>
	<li>After fixation, remove salivary glands, fat tissue and pull the brain away from the eye-antennal imaginal disc. To improve accessibility of 'difficult' antibodies, the peripodial membrane, which is located on the surface of the epithelium, can be removed by holding down the antennal part of the disc and peeling the membrane away with a dissecting pin. Next, perform immunohistochemistry (step 5.1.).</li>
</ol>
2. 50% Pupation/Midpupal Retinas
<ol>
	<li>Hold posterior end of the pupal case with a forceps. Gently remove the anterior surface of the pupal case by grabbing the cuticle with another forceps and slowly pull away from the puparium. Peel away residual cuticle to expose the head.</li>
	<li>Use the forceps' tips to pierce the underlying, soft sack in the dorsal anterior region. This provides access to the head. Gently separate the head from the thorax by slowly sucking up the head with a P20 pipette. Expel head from pipette tip into fresh cold 1x PBS and remove excess tissue; grab the head cuticle and remove proboscis. Proceed to fixation step (step 4.1.).</li>
	<li>Pierce the optic lobe with one dissecting pin to steady the tissue. Insert the other dissecting pin between the lamina and retina to separate the retina from the lamina/optic lobe. Perform immunohistochemistry (step 5.1.).</li>
</ol>
3. Adult Eyes
<ol>
	<li>Grab the center of the posterior head capsule with a forceps and remove mouth parts (proboscis), fat tissue and tracheae.</li>
	<li>Grab the dorsal head cuticle with one forceps and use the other to separate eyes from brain tissue by gently pulling sideways. In most cases, the lamina, a thin and transparent sheath of brain neuropil, will stay attached to the retina. It is easier to remove the lamina after fixation.</li>
	<li>Remove cuticle at the eye margin, but leave a small piece behind to facilitate the transfer of retinas to different solutions. Perform fixation (step 4.1.).</li>
	<li>Remove the lamina by grabbing it at one side with a forceps and gently pulling sideways with the other forceps without impairing the retina. Remove residual cuticle, as it could fold onto the retina during mounting. These two steps are critical for visualizing photoreceptors, which would otherwise be obscured by the lamina and remains of cuticle during the imaging process. Proceed to immunohistochemistry (step 5.1.).</li>
</ol>
4. Fixation and Washing (About 1 hr)
<ol>
	<li>Store the dissected retinas in 1x PBS in a three-well glass dish placed on ice while freshly preparing the fixative (dilute 40 microliters of 37% formaldehyde solution with 360 &mu;l of 1x PBS, vortex and store on ice). We recommend to proceed with the fixation protocol and to avoid long storage times on ice (&gt;15 min).</li>
	<li>Replace 1x PBS with 200 &mu;l of 3.7% formaldehyde solution. Make sure the tissue is submerged in fixative. This is critical, as it may otherwise not be fixed properly. If necessary, remove air bubbles and tracheae with the forceps.</li>
	<li>Incubate on shaker for 15 min at room temperature.</li>
	<li>Replace fixative with 1x PBS and rinse twice with 1x PBS. Continue washing in PBS for 30 min (on shaker, at room temperature).</li>
	<li>Replace PBS with PBST. Rinse twice with PBST.</li>
	<li>Proceed with removing the peripodial membrane from larval eye discs (3.1.3.) and the laminas from midpupal/adult retinas (3.2.3., 3.3.4.).</li>
</ol>
5. Immunohistochemistry
<ol>
	<li>Blocking: Replace PBS with 200 &mu;l of 5% normal serum in PBST and incubate retinas for 20 min on shaker at room temperature.</li>
	<li>Replace blocking solution with primary antibody solution. Seal the three-well dish with parafilm and incubate overnight on shaker at room temperature.</li>
	<li>Replace solution with PBST and rinse 3 times. Continue washing with PBST for a minimum of 1 hr on shaker at room temperature.</li>
	<li>Transfer in secondary antibody solution and incubate in a parafilm-sealed glass well dish on shaker overnight at room temperature.</li>
	<li>Replace antibody solution with PBST and rinse three times. Continue washing with PBST for a minimum of one hr on shaker at room temperature. The samples can be kept in PBST for several days, but it is critical to add fresh PBST at least once per day to prevent them from drying out.</li>
</ol>
6. Mounting
<ol>
	<li>For mounting larval and midpupal retinas, use the P20 to transfer retinas to the center of a clean glass slide. Remove excess PBST. Use a forceps to align retinas in order to facilitate imaging.</li>
	<li>Add 10 &mu;l of mounting medium on top of the retinas. Cover them gently with a 24x40-1 cover slip and seal with clear nail polish.</li>
	<li>For mounting adult retinas, prepare 'bridges' 20: Add two 5 &mu;l droplets of 50% glycerol on both ends of a glass side and cover them with 22x22-1 cover slips (0.13 to 0.17 mm thick).</li>
	<li>Remove the PBST with a P20 pipette; make sure that the retinas do not dry out. Add 20 &mu;l of mounting medium to the well. Use mounting medium to transfer retinas with the P20 to the 'bridge' glass slide.</li>
	<li>Use a forceps to orient (lenses should face the glass slide) and align retinas in order to facilitate imaging. If necessary, remove air bubbles with the P20.</li>
	<li>Slowly place 24x40-1 cover slip (0.13 to 0.17 mm thick) from one side on top and seal edges with clear nail polish or Scotch tape. Store samples at 4 &deg;C in a microscope slide saver box (keep in the dark).</li>
</ol>
7. Representative Results
As an example for the application of this protocol, we show representative results for visualizing photoreceptor differentiation at the three developmental stages in the video. A larval eye-antennal disc and a midpupal retina were stained with antibodies that label different photoreceptor types during their development (Figure 1A, B) and an adult retina was stained with two Rhodopsin antibodies to visualize two photoreceptor subtypes (Figure 1C).

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H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patterning+of+the+R7+and+R8+photoreceptor+cells+of+Drosophila:+evidence+for+induced+and+default+cell-fate+specification.">Patterning of the R7 and R8 photoreceptor cells of Drosophila: evidence for induced and default cell-fate specification.</a> Development. 126, 607-616 (1999).</li><li>Papatsenko, D., Sheng, G., Desplan, 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+new+rhodopsin+in+R8+photoreceptors+of+Drosophila:+evidence+for+coordinate+expression+with+Rh3+in+R7+cells.">A new rhodopsin in R8 photoreceptors of Drosophila: evidence for coordinate expression with Rh3 in R7 cells.</a> Development. 124, 1665-1673 (1997).</li><li>Wolff, T., Ready, D. 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No conflicts of interest declared.

1. Troubleshooting
In our experience, dissections require practice (up to several weeks) and are facilitated by achieving a comfortable hand position 21 by resting the elbows and forearms on the table and with the fingers making contact with the dissection dish. That way, only the thumbs, index and middle fingers perform subtle movements.
Removing the lamina without damaging the photoreceptors is probably the most challenging step. Practice with red-eyed wildtype flies first before dissecting white-eyed mutants, as the lamina is much easier to see against the red eye pigmentation. Also, use lighting from behind and a dark background below the dissection dish to achieve optimal contrast.
However, the red eye pigment is fluorescent and can cause high background during the imaging process. This can weaken the signals from the secondary antibodies. A good strategy is to keep washing the adult retinas for four to five days and to replace the washing solution with fresh PBST at least once per day. The pigment will be washed out; be sure to wash experimental and control genotypes for the same duration.
2. Applications
Eye-specific loss- and gain-of-function approaches 22 23 make the Drosophila eye a particularly powerful system for genetic manipulations and screens. Moreover, recently developed computational methods for single-cell analysis in 3D reconstructions of entire retinas allow the analysis of mutant phenotypes at high resolution 24. This protocol can therefore be applied to a variety of questions concerning eye development and the underlying regulatory mechanisms 16,25-28 .
<img alt="Figure 1" src="/files/ftp_upload/4347/4347fig1.jpg" /> 
Figure 1. Retinal whole mounts at three different developmental stages. A) Larval eye imaginal disc stained with antibodies against Seven-up (green) and Runt (magenta). B) Midpupal retina. Two antibodies were used to visualize all photoreceptors R1-R8 (Elav, blue) and R7 photoreceptors (Prospero, cyan). C) Adult retina. Stochastic and mutually exclusive expression of photosensitive pigments Rh5 (cyan) and Rh6 (red) in two subtypes of R8 photoreceptors.

<table border="1">
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Reagent</td>
 </tr>
 <tr>
 <td>Phosphate-buffered saline (PBS1x, pH 7.4)</td>
 <td>Sigma</td>
 <td></td>
 <td>Prepare 10x stock solution 20. Dilute with distilled water to obtain 1x PBS and store at room temperature. Cool on ice before dissections. </td>
 </tr>
 <tr>
 <td>Triton-X 100</td>
 <td>Sigma</td>
 <td>9002-93-1</td>
 <td>Caution: Irritant! Wear gloves. Prepare 50 ml 1xPBS with 0.3% Triton X-100 (PBST). Store at room temperature.</td>
 </tr>
 <tr>
 <td>37% formaldehyde solution </td>
 <td>Fisher Scientific</td>
 <td>F75P1GAL </td>
 <td>Caution: Toxic, probable human carcinogen! Wear gloves. Before the fixation step, freshly prepare 3.7% solution in a chemical fume hood by diluting with PBS, store on ice. </td>
 </tr>
 <tr>
 <td>5% normal horse serum </td>
 <td>Jackson Immuno Research</td>
 <td>008-000-001</td>
 <td>Prepare 5% v/v dilution in PBST. Store at four degrees.</td>
 </tr>
 <tr>
 <td>Primary and secondary antibodies (e.g. Donkey anti-sheep Alexa Fluor 488, Donkey-anti rabbit Alexa Fluor 555, Donkey anti-mouse Alexa Fluor 647) </td>
 <td>Invitrogen Molecular Probes</td>
 <td>A11015 A31572 A31571</td>
 <td>Dilute secondary antibodies 1:800 in PBST and store at four degrees. </td>
 </tr>
 <tr>
 <td>Alexa Fluor 488 Phalloidin</td>
 <td></td>
 <td>A12379 </td>
 <td>Dilute 1:100 in secondary antibody solution.</td>
 </tr>
 <tr>
 <td>Slowfade Gold Antifade reagent</td>
 <td>Invitrogen Molecular Probes</td>
 <td>S36936</td>
 <td>Mounting medium. Store at -20 degrees.</td>
 </tr>
 <tr>
 <td>Glycerol </td>
 <td>Fisher Scientific</td>
 <td>G31-1 </td>
 <td>For mounting. Prepare 10 ml of 50% dilution with distilled water, store at room temp.</td>
 </tr>
 <tr>
 <td>CO2</td>
 <td></td>
 <td></td>
 <td>For anesthetizing adult flies.</td>
 </tr>
 <tr>
 <td></td>
 <td></td>
 <td></td>
 <td>Equipment</td>
 </tr>
 <tr>
 <td>Two sharp forceps (Dumont #55)</td>
 <td>Fine Science Tools</td>
 <td>11255-20 </td>
 <td></td>
 </tr>
 <tr>
 <td>Sylgard 184 Silicone Elastomer Kit</td>
 <td>Dow Corning</td>
 <td></td>
 <td>Prepare Sylgard dissection dish by filling a plastic Petri dish with Sylgard mixture.</td>
 </tr>
 <tr>
 <td>Three-well glass dissection dishes</td>
 <td>Fisher Scientific</td>
 <td>21-379 </td>
 <td></td>
 </tr>
 <tr>
 <td>Two minutien dissecting pins (0.1 mm diameter) and two pinholders (12 cm)</td>
 <td>Fine Science tools </td>
 <td>26002-10</td>
 <td>Insert one minutien pin in each of the two pinholders and bend one of the pins to form a hook.</td>
 </tr>
 <tr>
 <td>Microscope cover slips (22x22-1 and 24x40-1)</td>
 <td>Fisher Scientific</td>
 <td>12-542A</td>
 <td></td>
 </tr>
 <tr>
 <td>Microscope slides (25x75x1.0 mm; precleaned)</td>
 <td>Fisher Scientific</td>
 <td>12-550-143</td>
 <td></td>
 </tr>
 <tr>
 <td>1.5 ml microcentrifuge tubes</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Clear nail polish or Scotch tape</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Orbital shaker</td>
 <td>Bellco</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Slide holder box</td>
 <td>Fisher Scientific</td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Parafilm</td>
 <td>Bemis</td>
 <td>PM996</td>
 <td></td>
 </tr>
 <tr>
 <td>Thin paintbrush </td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>P20 and P200 micropipettes and tips</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Dissecting microscope</td>
 <td></td>
 <td></td>
 <td></td>
 </tr>
 <tr>
 <td>Small bucket with ice</td>
 <td></td>
 <td></td>
 <td>For cooling the glass well plates, dissected retinas and solutions.</td>
 </tr>
 </table>

dissection and immunohistochemistry of larval, pupal and adult drosophila retinas

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including lens-secreting cone cells, pigment cells, a bristle cell and eight photoreceptors (PRs) R1-R8 2. The PRs have specialized microvillar structures, the rhabdomeres, which contain light-sensitive pigments, the Rhodopsins (Rhs). The rhabdomeres of six PRs (R1-R6) form a trapezoid and contain Rh1 3 4. The rhabdomeres of R7 and R8 are positioned in tandem in the center of the trapezoid and share the same path of light. R7 and R8 PRs stochastically express different combinations of Rhs in two main subtypes5: In the 'p' subtype, Rh3 in pR7s is coupled with Rh5 in pR8s, whereas in the 'y' subtype, Rh4 in yR7s is associated with Rh6 in yR8s 6 7 8.
Early specification of PRs and development of ommatidia begins in the larval eye-antennal imaginal disc, a monolayer of epithelial cells. A wave of differentiation sweeps across the disc9 and initiates the assembly of undifferentiated cells into ommatidia10-11. The 'founder cell' R8 is specified first and recruits R1-6 and then R7 12-14. Subsequently, during pupal development, PR differentiation leads to extensive morphological changes 15, including rhabdomere formation, synaptogenesis and eventually rh expression.
In this protocol, we describe methods for retinal dissections and immunohistochemistry at three defined periods of retina development, which can be applied to address a variety of questions concerning retinal formation and developmental pathways. Here, we use these methods to visualize the stepwise PR differentiation at the single-cell level in whole mount larval, midpupal and adult retinas (Figure 1).

Watch this Scientific Journal Video about Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas at JoVE.com

 New York University.  The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

25.1K Views.  New York University. The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

Video: Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The Drosophila retina is a crystal-like lattice composed of a small number of cell types that are generated in a stereotyped manner 1. Its amenability to sophisticated genetic analysis allows the study of complex developmental programs. This protocol describes dissections and immunohistochemistry of retinas at three discrete developmental stages, with a focus on photoreceptor differentiation.

Dissection and Immunohistochemistry of Larval, Pupal and Adult Drosophila Retinas

The compound eye of Drosophila melanogaster consists of about 750 ommatidia (unit eyes). Each ommatidium is composed of about 20 cells, including ...

dissection-immunohistochemistry-larval-pupal-adult-drosophila

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Neuroscience

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 expression is rarely tissue-specific enough to allow for precise correlation of the circuit with behavioral readouts. To overcome this major hurdle, we recently developed the FINGR method to achieve a more restrictive Gal4 expression in the tissue of interest. The FINGR method has three components: 1) the traditional Gal4/UAS system; 2) a set of FLP/FRT-mediated Gal80 converting tools; and 3) enhancer-trap FLP (ET-FLP). Gal4 is used to define the primary neural circuitry of interest. Paring the Gal4 with a UAS-effector, such as UAS-MJD78Q or UAS-Shits, regulates the neuronal activity, which is in turn manifested by alterations in the fly behavior. With an additional UAS-reporter such as UAS-GFP, the neural circuit involved in the specific behavior can be simultaneously mapped for morphological analysis. For Gal4 lines with broad expression, Gal4 expression can be restricted by using two complementary Gal80-converting tools: tubP&gt;Gal80&gt; ('flip out') and tubP&gt;stop&gt;Gal80 ('flip in'). Finally, investigators can turn Gal80 on or off, respectively, with the help of tissue-specific ET-FLP. In the flip-in mode, Gal80 will repress Gal4 expression wherever Gal4 and ET-FLP intersect. In the flip-out mode, Gal80 will relieve Gal4 repression in cells in which Gal4 and FLP overlap. Both approaches enable the restriction of the number of cells in the Gal4-defined circuitry, but in an inverse pattern. The FINGR method is compatible with the vast collection of Gal4 lines in the fly community and highly versatile for traditional clonal analysis and for neural circuit mapping. In this protocol, we demonstrate the mapping of FLP expression patterns in select ET-FLPx2 lines and the effectiveness of the FINGR method in photoreceptor cells. The principle of the FINGR method should also be applicable to other genetic model organisms in which Gal4/UAS, Gal80, and FLP/FRT are used.

Mapping and Application of Enhancer-trap Flippase Expression in Larval and Adult Drosophila CNS

We describe a Flippase-induced intersectional Gal80/Gal4 repression (FINGR) method, allowing tissue-specific FLP to determine Gal80 expression patterns. Wherever Gal4 and FLP overlap, Gal4 expression is turned on (Gal80 flipped out) or off (Gal80 flipped in). The FINGR method is versatile for clonal analysis and neural circuit mapping.

The Gal4/ UAS binary method is powerful for gene and neural circuitry manipulation in Drosophila. For most neurobiological studies, however, Gal4 ...

Single Drosophila Ommatidium Dissection and Imaging

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative diseases because of its powerful genetics1. The fly eye in particular has been the organ of choice for neurodegeneration research, being the most accessible and life-dispensable part of the Drosophila nervous system. However the major caveat of intact eyes is the difficulty, because of the intense autofluorescence of the pigment, in imaging intracellular events, such as autophagy dynamics2, which are paramount to understanding of neurodegeneration.
We have recently used the dissection and culture of single ommatidia3 that has been essential for our understanding of autophagic dysfunctions in a fly model of Dentatorubro-Pallidoluysian Atrophy (DRPLA)3, 4.
We now report a comprehensive description of this technique (Fig. 1), adapted from electrophysiological studies5, which is likely to expand dramatically the possibility of fly models for neurodegeneration. This method can be adapted to image live subcellular events and to monitor effective drug administration onto photoreceptor cells (Fig. 2). If used in combination with mosaic techniques6-8, the responses of genetically different cells can be assayed in parallel (Fig. 2).

Single Drosophila Ommatidium Dissection and Imaging

The limiting factor in the use of the adult Drosophila eye to study neurodegeneration and cell biology is the difficult imaging of intracellular processes. We describe the dissection of single ommatidia to generate a bona-fide primary neuronal cell culture, which can be subject to drug treatment and advanced imaging.

The fruit fly Drosophila melanogaster has made invaluable contributions to neuroscience research and has been used widely as a model for neurodegenerative ...

Dissection of Adult Mouse Utricle and Adenovirus-mediated Supporting-cell Infection

Hearing loss and balance disturbances are often caused by death of mechanosensory hair cells, which are the receptor cells of the inner ear. Since there is no cell line that satisfactorily represents mammalian hair cells, research on hair cells relies on primary organ cultures. The best-characterized in vitro model system of mature mammalian hair cells utilizes organ cultures of utricles from adult mice (Figure 1) 1-6. The utricle is a vestibular organ, and the hair cells of the utricle are similar in both structure and function to the hair cells in the auditory organ, the organ of Corti. The adult mouse utricle preparation represents a mature sensory epithelium for studies of the molecular signals that regulate the survival, homeostasis, and death of these cells.
Mammalian cochlear hair cells are terminally differentiated and are not regenerated when they are lost. In non-mammalian vertebrates, auditory or vestibular hair cell death is followed by robust regeneration which restores hearing and balance functions 7, 8. Hair cell regeneration is mediated by glia-like supporting cells, which contact the basolateral surfaces of hair cells in the sensory epithelium 9, 10. Supporting cells are also important mediators of hair cell survival and death 11. We have recently developed a technique for infection of supporting cells in cultured utricles using adenovirus. Using adenovirus type 5 (dE1/E3) to deliver a transgene containing GFP under the control of the CMV promoter, we find that adenovirus specifically and efficiently infects supporting cells. Supporting cell infection efficiency is approximately 25-50%, and hair cells are not infected (Figure 2). Importantly, we find that adenoviral infection of supporting cells does not result in toxicity to hair cells or supporting cells, as cell counts in Ad-GFP infected utricles are equivalent to those in non-infected utricles (Figure 3). Thus adenovirus-mediated gene expression in supporting cells of cultured utricles provides a powerful tool to study the roles of supporting cells as mediators of hair cell survival, death, and regeneration.

Mechanosensory hair cells are the receptor cells of the inner ear. The best-characterized in vitro model system of mature mammalian hair cells utilizes organ cultures of utricles from adult mice. We present the dissection of the adult mouse utricle, and we demonstrate adenovirus-mediated infection of supporting cells in cultured utricles.

Hearing loss and balance disturbances are often caused by death of mechanosensory hair cells, which are the receptor cells of the inner ear.  Since there ...

Functional Analysis of the Larval Feeding Circuit in Drosophila

The serotonergic feeding circuit in Drosophila melanogaster larvae can be used to investigate neuronal substrates of critical importance during the development of the circuit. Using the functional output of the circuit, feeding, changes in the neuronal architecture of the stomatogastric system can be visualized. Feeding behavior can be recorded by observing the rate of retraction of the mouth hooks, which receive innervation from the brain. Locomotor behavior is used as a physiological control for feeding, since larvae use their mouth hooks to traverse across an agar substrate. Changes in feeding behavior can be correlated with the axonal architecture of the neurites innervating the gut. Using immunohistochemistry it is possible to visualize and quantitate these changes. Improper handling of the larvae during behavior paradigms can alter data as they are very sensitive to manipulations. Proper imaging of the neurite architecture innervating the gut is critical for precise quantitation of number and size of varicosities as well as the extent of branch nodes. Analysis of most circuits allow only for visualization of neurite architecture or behavioral effects; however, this model allows one to correlate the functional output of the circuit with the impairments in neuronal architecture.

Functional Analysis of the Larval Feeding Circuit in Drosophila

The feeding circuit in Drosophila melanogaster larvae serves a simple yet powerful model that allows changes in feeding rate to be correlated with alterations in the stomatogastric neural circuitry. This circuit is composed of central serotonergic neurons that send projections to the mouth hooks as well as the foregut.

The serotonergic  feeding circuit in Drosophila  melanogaster larvae can be used to investigate neuronal substrates of  critical importance during the ...

Live Imaging of Drosophila Larval Neuroblasts

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given their relevance to development and disease, understanding the mechanisms that govern asymmetric stem cell division has been a robust area of study. Because they are genetically tractable and undergo successive rounds of cell division about once every hour, the stem cells of the Drosophila central nervous system, or neuroblasts, are indispensable models for the study of stem cell division. About 100 neural stem cells are located near the surface of each of the two larval brain lobes, making this model system particularly useful for live imaging microscopy studies. In this work, we review several approaches widely used to visualize stem cell divisions, and we address the relative advantages and disadvantages of those techniques that employ dissociated versus intact brain tissues. We also detail our simplified protocol used to explant whole brains from third instar larvae for live cell imaging and fixed analysis applications.

Live Imaging of Drosophila Larval Neuroblasts

This protocol details a streamlined method used to conduct live cell imaging in the context of an intact larval brain. Live cell imaging approaches are invaluable for the study of asymmetric neural stem cell divisions as well as other neurogenic and developmental processes, consistently uncovering mechanisms that were previously overlooked.

Stem cells divide asymmetrically to generate two progeny cells with unequal fate potential: a self-renewing stem cell and a differentiating cell. Given ...

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

Drosophila melanogaster is a powerful experimental model system for studying the function of the nervous system. Gene mutations that cause dysfunction of the nervous system often produce viable larvae and adults that have locomotion defective phenotypes that are difficult to adequately describe with text or completely represent with a single photographic image. Current modes of scientific publishing, however, support the submission of digital video media as supplemental material to accompany a manuscript. Here we describe a simple and widely accessible microscopy technique for acquiring high-quality digital video of both Drosophila larval and adult phenotypes from a lateral perspective. Video of larval and adult locomotion from a side-view is advantageous because it allows the observation and analysis of subtle distinctions and variations in aberrant locomotive behaviors. We have successfully used the technique to visualize and quantify aberrant crawling behaviors in third instar larvae, in addition to adult mutant phenotypes and behaviors including grooming.

Acquisition of High-Quality Digital Video of Drosophila Larval and Adult Behaviors from a Lateral Perspective

Here we describe a simple and widely accessible microscopy technique to acquire high-quality digital video of Drosophila adult and larval mutant phenotypes from a lateral perspective.

Drosophila melanogaster is a powerful experimental model system for studying the function of the nervous system. Gene mutations that cause dysfunction of ...

Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a spiral shape and have a tonotopic gradient for sound detection. The mouse cochlea is approximately 6 mm long and often divided into three turns (apex, middle, and base) for analysis. To investigate cell loss, cell division, or mosaic gene expression, the whole mount or surface preparation of the cochlea is useful. This dissection method allows visualization of all cells within the organ of Corti when combined with immunostaining and confocal microscopy to image cells at different planes in the z-axis. Multiple optical cross-sections can also be obtained from these z-stack images. In addition, the whole mount dissection method can be used for scanning electron microscopy, although a different fixation method is needed. Here, we present a method to isolate the organ of Corti as three intact cochlear turns (apex, middle, and base). This method can be used for mice ranging from one week of age through adulthood and differs from the technique used for neonatal samples where calcification of the cochlea is incomplete. A slightly modified version can be used for dissection of the rat cochlea. We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies.

We present a method to isolate the adult organ of Corti as three intact cochlear turns (apex, middle, and base). We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies. Together these techniques allow the study of hair cells, supporting cells, and other cell types found in the cochlea.

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a ...

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat open surfaces is complex,&#160;including: pausing, turning, and meandering. This complexity in the repertoire of movement hinders detailed analysis of the events occurring during a single crawl stride cycle. To overcome this obstacle, linear agarose channels were made that constrain larval behavior to straight, sustained, rhythmic crawling. In principle, because agarose channels and the Drosophila larval body are both optically clear, the movement of larval structures labeled by genetically-encoded fluorescent probes can be monitored in intact, freely-moving larvae. In the past, larvae were placed in linear channels and crawling at the level of whole organism, segment, and muscle were analyzed1. In the future, larvae crawling in channels can be used for calcium imaging to monitor neuronal activity. Moreover, these methods can be used with larvae of any genotype and with any researcher-designed channel. Thus the protocol presented below is widely applicable for studies using the Drosophila larva as a model to understand motor control.

Using Linear Agarose Channels to Study Drosophila Larval Crawling Behavior

The Drosophila larva is a powerful model system to study neural control of behavior. This publication describes the use of linear agarose channels to elicit sustained bouts of linear crawling and methods to quantify the dynamics of larval structures during repetitive crawling behavior.

Drosophila larval crawling is emerging as a powerful model to study neural control of sensorimotor behavior. However, larval crawling behavior on flat ...

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the molecular and cellular basis of higher brain functions. A whole-mount preparation of adult brains with well-preserved morphology is critical for such whole brain-based studies, but can be technically challenging and time-consuming. This protocol describes an easy-to-learn, one-step dissection approach of an adult fly head in less than 10 s, while keeping the intact brain attached to the rest of the body to facilitate subsequent processing steps. The procedure helps remove most of the eye and tracheal tissues normally associated with the brain that can interfere with the later imaging step, and also places less demand on the quality of the dissecting forceps. Additionally, we describe a simple method that allows convenient flipping of the mounted brain samples on a coverslip, which is important for imaging both sides of the brains with similar signal intensity and quality. As an example of the protocol, we present an analysis of dopaminergic (DA) neurons in adult brains of WT (w1118) flies. The high efficacy of the dissection method makes it particularly useful for large-scale adult brain-based studies in Drosophila.

A Simple One-step Dissection Protocol for Whole-mount Preparation of Adult Drosophila Brains

The adult Drosophila brain is a valuable system for studying neuronal circuitry, higher brain functions, and complex disorders. An efficient method to dissect whole brain tissue from the small fly head will facilitate brain-based studies. Here we describe a simple, one-step dissection protocol of adult brains with well-preserved morphology.

There is an increasing interest in using Drosophila to model human brain degenerative diseases, map neuronal circuitries in adult brains, and study the ...

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs provides a tissue-level understanding of energy utilization that cannot be captured when analyzing primary cells and cell lines. While this analysis is ex vivo, it allows for the measurement from a number of specialized cells working together to perform a function in one tissue and more closely models the in vivo organ. Metabolic reprogramming has been observed in many neurological diseases, including neoplasia, and neurodegenerative diseases. This protocol was developed to assist the D. melanogaster community&#39;s investigation of metabolism in neurological disease models using a commercially available metabolic analyzer. Measuring metabolism of whole brains in the metabolic analyzer is challenging due to the geometry of the brain. This analyzer requires samples to remain at the bottom of a 96-well plate. Cell samples and tissue punches can adhere to the surface of the cell plate or utilize spheroid plates, respectively. However, the spherical, three-dimensional shape of D. melanogaster brains prevents the tissue from adhering to the plate. This protocol requires a specially designed and manufactured micro-tissue restraint that circumvents this problem by preventing any movement of the brain while still allowing metabolic measurements from the analyzer&#39;s two solid-state sensor probes. Oxygen consumption and extracellular acidification rates are reproducible and sensitive to a treatment with metabolic inhibitors. With a minor optimization, this protocol can be adapted for use with any whole tissue and/or model system, provided that the sample size does not exceed the chamber generated by the restraint. While basal metabolic measurements and an analysis after a treatment with mitochondrial inhibitors are described within this protocol, countless experimental conditions, such as energy source preference and rearing environment, could be interrogated.

Metabolic Analysis of Drosophila melanogaster Larval and Adult Brains

We present a protocol for measuring oxygen consumption and extracellular acidification in Drosophila melanogaster larval and adult brains. A metabolic analyzer is utilized with an adapted and optimized protocol. Micro-tissue restraints are a critical component of this protocol and were designed and created specifically for their use in this analysis.

This protocol describes a method for measuring the metabolism in Drosophila melanogaster larval and adult brains. Quantifying metabolism in whole organs ...