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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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 w...

<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).

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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

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

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of its convenient genetics and fully sequenced genome. Additionally, the larval nervous system is an ideal model system to study mechanisms of axonal transport as the larval segmental nerves contain bundles of axons with their cell bodies located within the brain and their nerve terminals ending along the length of the body. Here we describe the procedure for visualization of synaptic vesicle proteins within larval segmental nerves. If done correctly, all components of the nervous system, along with associated tissues such as muscles and NMJs, remain intact, undamaged, and ready to be visualized. 3rd instar larvae carrying various mutations are dissected, fixed, incubated with synaptic vesicle antibodies, visualized and compared to wild type larvae. This procedure can be adapted for several different synaptic or neuronal antibodies and changes in the distribution of a variety of proteins can be easily observed within larval segmental nerves.

Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations

Drosophila melanogaster larvae provide an ideal model system to investigate the mechanisms of axonal transport within larval segmental nerves. Using this procedure, 3rd instar larvae carrying various mutations can be compared to wild type larvae.

Drosophila melanogaster is emerging as a powerful model system for studying the development and function of the nervous system, particularly because of ...

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 ...

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

The Drosophila larvae neuromuscular junction (NMJ) is an excellent model for the study of synaptic structure and function. Drosophila is well known for the ease of powerful genetic manipulations and the larval nervous system has proven particularly useful in studying not only normal function but also perturbations that accompany some neurological disease (Lloyd and Taylor, 2010). Many key synaptic molecules found in Drosophila are also found in mammals and like most CNS excitatory synapses in mammals, the Drosophila NMJ is glutamatergic and demonstrates activity-dependent remodeling (Kohet al. , 2000). Additionally, Drosophila neurons can be individually identified because their innervation patterns are stereotyped and repetitive making it possible to study identified synaptic terminals, such as those between motor neurons and the body-wall muscle fibers that they innervate (Keshishian and Kim, 2004). The existence of evolutionarily conserved synapse components along with the ease of genetic and physical manipulation make the Drosophila model ideal for investigating the mechanisms underlying synaptic function (Budnik, 1996).
The active zones at synaptic terminals are of particular interest because these are the sites of neurotransmitter release. NC82 is a monoclonal antibody that recognizes the Drosophila protein Bruchpilot (Brp), a CAST1/ERC family member that is an important component of the active zone (Waghet al. , 2006). Brp was shown to directly shape the active zone T-bar and is responsible for effectively clustering Ca2+ channels beneath the T-bar density (Fouquetet al. , 2009). Mutants of Brp have reduced Ca2+ channel density, depressed evoked vesicle release, and altered short-term plasticity (Kittelet al. , 2006). Alterations to active zones have been observed in Drosophila disease models. For example, immunofluorescence using the NC82 antibody showed that the active zone density was decreased in models of amyotrophic lateral sclerosis and Pitt-Hopkins syndrome (Ratnaparkhiet al. , 2008; Zweieret al. , 2009). Thus, evaluation of active zones, or other synaptic proteins, in Drosophila larvae models of disease may provide a valuable initial clue to the presence of a synaptic defect.
Preparing whole-mount dissected Drosophila larvae for immunofluorescence analysis of the NMJ requires some skill, but can be accomplished by most scientists with a little practice. Presented is a method that provides for multiple larvae to be dissected and immunostained in the same dissection dish, limiting environmental differences between each genotype and providing sufficient animals for confidence in reproducibility and statistical analysis.

Dissection and Imaging of Active Zones in the Drosophila Neuromuscular Junction

The neuromuscular junction (NMJ) of Drosophila melanogaster is an important model system for studying normal synaptic function as well as perturbations to synaptic function found in certain neurological diseases. We present a protocol for dissection of the Drosophila larval motor system and immunostaining for active zone proteins within the NMJ.

The Drosophila  larvae neuromuscular junction (NMJ) is an excellent model for the study of synaptic structure and function. Drosophila  is well known for ...

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 ...

Drosophila Adult Olfactory Shock Learning

Drosophila have been used in classical conditioning experiments for over 40 years, thus greatly facilitating our understanding of memory, including the elucidation of the molecular mechanisms involved in cognitive diseases1-7. Learning and memory can be assayed in larvae to study the effect of neurodevelopmental genes8-10 and in flies to measure the contribution of adult plasticity genes1-7. Furthermore, the short lifespan of Drosophila facilitates the analysis of genes mediating age-related memory impairment5,11-13. The availability of many inducible promoters that subdivide the Drosophila nervous system makes it possible to determine when and where a gene of interest is required for normal memory as well as relay of different aspects of the reinforcement signal3,4,14,16.
Studying memory in adult Drosophila allows for a detailed analysis of the behavior and circuitry involved and a measurement of long-term memory15-17. The length of the adult stage accommodates longer-term genetic, behavioral, dietary and pharmacological manipulations of memory, in addition to determining the effect of aging and neurodegenerative disease on memory3-6,11-13,15-21.
Classical conditioning is induced by the simultaneous presentation of a neutral odor cue (conditioned stimulus, CS+) and a reinforcement stimulus, e.g., an electric shock or sucrose, (unconditioned stimulus, US), that become associated with one another by the animal1,16. A second conditioned stimulus (CS-) is subsequently presented without the US. During the testing phase, Drosophila are simultaneously presented with CS+ and CS- odors. After the Drosophila are provided time to choose between the odors, the distribution of the animals is recorded. This procedure allows associative aversive or appetitive conditioning to be reliably measured without a bias introduced by the innate preference for either of the conditioned stimuli. Various control experiments are also performed to test whether all genotypes respond normally to odor and reinforcement alone.

Drosophila Adult Olfactory Shock Learning

The method to measure adult Drosophila associative memory is described. The assay is based on the ability of the fly to associate an odor presented with a negative reinforcer (electric shock) and then recall this information at a later time, allowing memory to be measured.

Drosophila have been used in classical conditioning experiments for over 40 years, thus greatly facilitating our understanding of memory, including the ...

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 ...