The authors thank RIKEN for funding. We also thank Cagri Yalgin, Caroline Delandre, and Jay Parrish for discussions on genetic and immunohistochemistry protocols.

1.Preparation of reagents
<ol>
 <li>Prepare Ca++ -free HL3.1 saline28.
 <ol>
 <li>In mM: 70 NaCl, 5 KCl, 20 MgCl2 , 10 NaHCO3 , 5 HEPES, 115 sucrose, and 5 trehalose; pH 7.2. Filter sterilize and store at 4&deg;C. 
 Note: Ca++ -free solution prevents muscle contraction during dissection.</li>
 </ol>
 </li>
 <li>Make poly-L-lysine (PLL) coverslips.
 <ol>
 <li>Dissolve 100mg PLL in 4.2ml water and make 300&mu;l aliquots in Eppendorf tubes and freeze at -20&deg;C.</li>
 <li>Before coating coverslips, first thaw an aliquot, bring it up to 10ml in ddH2O and add 20&mu;l of Kodak Photo-Flo; the final PLL concentration is 0.7 mg/ml.</li>
 <li>Submerge coverslips for 30mins in PLL solution, remove and dry, then rinse briefly with water and dry. Repeat twice. Treated coverslips last approximately one month.</li>
 </ol>
 </li>
</ol>
2. Genetic crosses
<ol>
 <li>To generate MARCM clones. Here is an example cross using a pan-DA driver11,27: 
 FRT2A x hsFLP; Gal4109(2)80, UAS-mCD8::GFP; tub-Gal80 FRT2A/SM5-TM6B</li>
 <li>To generate Flp-out clones. Here is an example cross using a class IV-specific driver8: 
 ppk-Gal4 x yw, hsFLP;UAS-FRT-CD2,y+-stop-FRT-mCD8::GFP</li>
</ol>
3. Collection of embryos
<ol>
 <li>Keep Drosophila crosses in a collection bottle at 25&deg;C and collect embryos on an apple juice agar plate29 spread with a thin layer of yeast paste30,31.</li>
</ol>
4. Heat shock treatment of embryos
<ol>
 <li>Place an empty plate in opposition to the apple juice agar plate upon which the embryos have been laid. Seal around these two plates with Parafilm (Fig. 2).</li>
 <li>During heat shock, submerge the plate in the water bath and hold it down using a metal weight.</li>
 <li>For MARCM embryos
 <ol>
 <li>Collect embryos for 2h and incubate in a 10cm petri dish surrounded by moistened tissues at 25&deg;C for an additional 2h. 
 Note: Adjusting the heat shock protocol alters the frequency at which clones are generated.</li>
 <li>For a smaller number of clones, aiming for single isolated neurons, submerge and heat shock in the water bath for 1h at 38&deg;C.</li>
 <li>For a larger number of clones, heat shock for 45mins at 38&deg;C, recover at RT 30mins, then heat shock again for an additional 30mins.</li>
 </ol>
 </li>
 <li>Collect Flp-out embryos for 24h, and heat shock for 1h at 38&deg;C.</li>
</ol>
5. Screening for clones
<ol>
 <li>After heat shock, remove the lid of the water-tight arrangement and place the apple juice agar plate in a 10cm petri dish surrounded by moistened tissues. Culture the embryos and subsequent larvae at 25&deg;C until wandering 3rd instar. 
 Note: culture conditions, especially nutrition, have been shown to alter DA dendritic arbor morphology32,33. Make sure that growing larvae have access to yeast paste at all times; monitor and replenish the yeast paste as required.</li>
 <li>From this point in the protocol onwards, manipulate larvae gently using insect forceps.</li>
 <li>Briefly and gently rinse the larvae in tap water, and then place onto an agar plate. 
 Note: This step reduces background auto-fluorescence from food or dirt on the larval surface.</li>
 <li>Examine the larvae under a powerful fluorescence dissecting microscope. Identify larvae with GFP-positive neurons/cells in the body wall.</li>
</ol>
6. in vivo imaging of dendrites
<ol>
 <li>Place the larva into a depression slide glass with a small drop of 80% glycerol. Place a coverslip on slide to immobilize the larva. Ensure that no air remains between the larva and the coverslip.</li>
 <li>Gently push the coverslip to roll the larva to allow visualization of the neuron of interest. Image the mCD8::GFP-labeled dendritic arbor via confocal microscopy.</li>
</ol>
7. Larval dissection
Note before beginning: DA neuron dendrites degrade rapidly after the initiation of dissection. Dissect each individual larva in less than 5min to ensure good dendrite morphology.
<ol>
 <li>Dissect wandering 3rd instar larvae on a Sylgard plate as described previously34 with the following modifications:
 <ol>
 <li>For the imaging of DA neuron axon termini, ensure that the CNS is intact and the segmental nerves are not broken. After opening the larvae, first carefully cut the tracheal connections to the body wall and then gently remove the entire gut.</li>
 <li>If imaging the dendrites alone, remove the CNS to allow a flatter mounting.</li>
 </ol>
 </li>
</ol>
8. Fixation and blocking of larval fillets
<ol>
 <li>With the larva still pinned to the Sylgard plate, fix it in 4% PFA in PBS for 20mins at RT on a gently rotating shaker.</li>
 <li>Remove traces of larval tissues (fat body, trachea, etc.) after fixation.</li>
 <li>Wash in PBST (PBS with 0.1% Triton X-100) 3x 10mins on the Sylgard plate and shaker. If examining axon termini, keep the fillet on the Sylgard plate. (If staining dendrites it is possible to transfer the larval fillets to a 0.5 ml tube at this step34)</li>
 <li>Block the larvae for 20mins at RT in 5% normal donkey serum (NDS) in PBST on a shaker.</li>
</ol>
9. Staining of larval fillets
<ol>
 <li>Remove the blocking solution and incubate in primary antibody (in 5%NDS/PBST) overnight at 4&deg;C in a small Tupperware container surrounded by moistened tissues.</li>
 <li>Remove the tubes from 4&deg;C and incubate an additional hour at RT.</li>
 <li>Wash 6x 10mins in PBST. Add the secondary antibody (in 5%NDS/PBST), and cover to prevent fluorophore photo-bleaching34.</li>
 <li>Incubate either at RT for two hours, or overnight at 4&deg;C followed by one hour at RT.</li>
 <li>Wash the larvae 6x 10mins in PBST and proceed to mounting.</li>
</ol>
10. Mounting of larval fillets for examination of the dendrite arbor
<ol>
 <li>Mount each larval fillet as flat as possible using dissecting scissors or a scalpel to cut off the head (including the mouth hooks), and the posterior (including the spiracles)34.</li>
 <li>Place the larval fillet on the slide cuticle-side down, mount in 80% glycerol, and seal the sides of the coverslip with nail polish for a 'quick' mount34.</li>
 <li>For a permanent mounting and a clearer image
 <ol>
 <li>Place the dissected larval fillet muscle-side down onto a drop of PBS on a PLL coverslip, it will quickly adhere to the coverslip.</li>
 <li>Remove as much liquid as possible after mounting, however do not allow it to completely dry.</li>
 <li>Take the coverslip (with attached larval fillet) through an ethanol series: 35% ethanol, followed by 50%, 70%, 95%, and finally 2x 100% ethanol each for 10 minutes (the 2nd 100% EtOH solution should be changed frequently); finally, wash 2-3x 10mins in Xylene.</li>
 <li>Put a drop of DPX mountant (Distyrene Plasticizer Xylene) on a clean slide and carefully lay the coverslip fillet-side-down on top. Keep in the dark, and wait one day for the DPX to set before imaging.</li>
 </ol>
 </li>
</ol>
11. Mounting of larval fillets for examination of the axon termini
<ol>
 <li>Keep the segmental nerves running between the PNS and the CNS intact during dissection, fixation, and staining to allow the tracing of the axons from peripheral sensory neuron cell body to the VNC.</li>
 <li>Mount each larval fillet cuticle-side down in 80% glycerol in a depression slide. Trace the axons from the DA cell body to the VNC under the confocal microscope. Note: PNS neurons in each body wall segment project to the cognate segment of the VNC.</li>
</ol>
12. Representative Results:
Representative results are shown in figures 3-5. Fig. 3 shows the entire arbor of a class IV da neuron, captured in vivo under the confocal microscope. Fig. 4 shows a close up of part of the dendrite arbor of an immunohistochemically labeled class III neuron that has been correctly fixed to preserve morphology. The associated inset shows the degradation that can occur after an unsuccessful dissection and fixation (both stained with anti-GFP antibody). Fig. 5 shows a single class IV axon terminus in the CNS (anti-GFP, green); all class IV termini are co-stained with anti-CD2 (magenta).
<img alt="Figure 1" src="/files/ftp_upload/3111/3111fig1.jpg" /> 
Figure 1 Protocol overview. a) Collect and then heat-shock embryos. b) Select a larva with a GFP-positive DA neuron clone, and then image the GFP-labeled dendrite arbor in the live larva. c) Dissect the larva, and then immunohistochemically stain the fillet. d) Mount the stained fillet, and then image the dendritic arbor and axonal projections of the DA neuron clones.
<img alt="Figure 2" src="/files/ftp_upload/3111/3111fig2.jpg" /> 
Figure 2 Preparation of the apple juice agar plate for heat shock. Take the plate (white arrow, a-b), add a second petri dish on top (red arrow b) and seal around with Parafilm (blue arrow, b).
<img alt="Figure 3" src="/files/ftp_upload/3111/3111fig3.jpg" /> 
Figure 3 in vivo image of the dendrite arbor of a mCD8::GFP-labeled MARCM clone (genotype as in 2.1) representing a class IV (v'ada) neuron of a 3rd instar larva. Scale bar is 50&mu;m.
<img alt="Figure 4" src="/files/ftp_upload/3111/3111fig4.jpg" /> 
Figure 4 mCD8::GFP-labeled MARCM clone of a class III neuron (ddaA) stained with anti-GFP antibody, showing good morphology after successful dissection and fixation (green). The inset shows dendrite degradation (white arrows) occurring after unsuccessful dissection and fixation (magenta). Scale bar is 25&mu;m.
<img alt="Figure 5" src="/files/ftp_upload/3111/3111fig5.jpg" /> 
Figure 5 The 3rd instar larval VNC. A Flp-out clone of a single class VI (vdaB) axon terminus (genotype as in 2.2) is detected using anti-GFP antibody (green). Anti-CD2 labels all class IV termini (magenta). Scale bar is 25&mu;m.

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Watching+a+synapse+grow:+noninvasive+confocal+imaging+of+synaptic+growth+in+Drosophila.">Watching a synapse grow: noninvasive confocal imaging of synaptic growth in Drosophila.</a> Neuron. 22, 719-729 (1999).</li><li>Landgraf, M., Sanchez-Soriano, N., Technau, G. M., Urban, J., Prokop, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Charting+the+Drosophila+neuropile:+a+strategy+for+the+standardised+characterisation+of+genetically+amenable+neurites.">Charting the Drosophila neuropile: a strategy for the standardised characterisation of genetically amenable neurites.</a> Dev Biol. 260, 207-225 (2003).</li></ol>

No conflicts of interest declared.

The Drosophila larval DA neuron model provides one excellent genetic system to investigate mechanisms that control neuron morphology and circuit formation. MARCM is generally used for labeling and for generating mutant DA neuron clones. For MARCM we use either a pan-neural (e.g. Gal4c155) or DA neuron-specific driver. Using a pan-neural driver it is possible to directly use several stocks widely available from public stock centers. However using a DA-neuron specific driver can be adv...

<table border="1">
<tr>
<th>Name of the reagent</th>
<th>Company</th>
<th>Catalogue number</th>
<th>Comments (optional)</th>
</tr>
<tr>
<td>SZX16 fluorescence dissection microscope (with GFPHQ filter)</td>
<td>Olympus</td>
<td>SZX16</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Live Insect Forceps</td>
<td>FST</td>
<td>26030-10</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>26mm x 76mm depression slide glass</td>
<td>Toshinriko Co.</td>
<td>T8-R004</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Sylgard 184 (or Silpot 184)</td>
<td>Dow Corning</td>
<td>3097358-1004</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Poly-L-lysine</td>
<td>Sigma </td>
<td>P-1524</td>
<td>This product has proven most effective</td>
</tr>
<tr>
<td>DPX mounting medium</td>
<td>Sigma</td>
<td>44581</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Rabbit anti-GFP</td>
<td>Invitrogen</td>
<td>A-11122</td>
<td>Dilution 1:500</td>
</tr>
<tr>
<td>Rat anti-CD8</td>
<td>Caltag </td>
<td>5H10</td>
<td>Dilution 1:200</td>
</tr>
<tr>
<td>Mouse anti-CD2</td>
<td>AbD serotec</td>
<td>MCA443R</td>
<td>Dilution 1:700</td>
</tr>
<tr>
<td>Mouse anti-Fasciclin2</td>
<td>DSHB</td>
<td>1D4</td>
<td>Dilution 1:10</td>
</tr>
</table>

morphological analysis of drosophila larval peripheral sensory neuron dendrites and axons using genetic mosaics

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic development and axonal wiring. Recently the dendritic arborization (DA) sensory neurons of the Drosophila larval peripheral nervous system (PNS) have become powerful genetic models in which to elucidate both general and class-specific mechanisms of neuron differentiation.
There are four main DA neuron classes (I-IV)1. They are named in order of increasing dendrite arbor complexity, and have class-specific differences in the genetic control of their differentiation2-10. The DA sensory system is a practical model to investigate the molecular mechanisms behind the control of dendritic morphology11-13 because: 1) it can take advantage of the powerful genetic tools available in the fruit fly, 2) the DA neuron dendrite arbor spreads out in only 2 dimensions beneath an optically clear larval cuticle making it easy to visualize with high resolution in vivo, 3) the class-specific diversity in dendritic morphology facilitates a comparative analysis to find key elements controlling the formation of simple vs. highly branched dendritic trees, and 4) dendritic arbor stereotypical shapes of different DA neurons facilitate morphometric statistical analyses.
DA neuron activity modifies the output of a larval locomotion central pattern generator14-16. The different DA neuron classes have distinct sensory modalities, and their activation elicits different behavioral responses14,16-20. Furthermore different classes send axonal projections stereotypically into the Drosophila larval central nervous system in the ventral nerve cord (VNC)21. These projections terminate with topographic representations of both DA neuron sensory modality and the position in the body wall of the dendritic field7,22,23. Hence examination of DA axonal projections can be used to elucidate mechanisms underlying topographic mapping7,22,23, as well as the wiring of a simple circuit modulating larval locomotion14-17.
We present here a practical guide to generate and analyze genetic mosaics24 marking DA neurons via MARCM (Mosaic Analysis with a Repressible Cell Marker)1,10,25 and Flp-out22,26,27 techniques (summarized in Fig. 1).

Watch this Scientific Journal Video about Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics at JoVE.com

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

The dendritic arborization sensory neurons of the Drosophila larval peripheral nervous system are useful models to elucidate both general and neuron class-specific mechanisms of neuron differentiation. We present a practical guide to generate and analyze dendritic arborization neuron genetic mosaics.

Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

Nervous system development requires the correct specification of neuron position and identity, followed by accurate neuron class-specific dendritic ...

morphological-analysis-drosophila-larval-peripheral-sensory-neuron

Research

JoVE Journal

Neuroscience

15.2K Aufrufe.  RIKEN Brain Science Institute. Die dendritischen Verzweigung sensorischen Neuronen des Drosophila Larven des peripheren Nervensystems sind nützliche Modelle, um sowohl allgemeine als auch Neuron-Klasse-spezifische Mechanismen der neuronalen Differenzierung aufzuklären. Wir präsentieren eine praktische Anleitung zur Generierung und Analyse dendritischer Verzweigung Neuron genetischen Mosaiken.

Serie: Morphological Analysis of Drosophila Larval Peripheral Sensory Neuron Dendrites and Axons Using Genetic Mosaics

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different neurological diseases. Defects in this essential process can have detrimental effects on neuronal functioning and development. We have developed a dissection protocol that is designed to expose the Drosophila larval segmental nerves to view axonal transport in real time. We have adapted this protocol for live imaging from the one published by Hurd and Saxton (1996) used for immunolocalizatin of larval segmental nerves. Careful dissection and proper buffer conditions are critical for maximizing the lifespan of the dissected larvae. When properly done, dissected larvae have shown robust vesicle transport for 2-3 hours under physiological conditions. We use the UAS-GAL4 method 1 to express GFP-tagged APP or synaptotagmin vesicles within a single axon or many axons in larval segmental nerves by using different neuronal GAL4 drivers. Other fluorescently tagged markers, for example mitochrondria (MitoTracker) or lysosomes (LysoTracker), can be also applied to the larvae before viewing. GFP-vesicle movement and particle movement can be viewed simultaneously using separate wavelengths.

In vivo Visualization of Synaptic Vesicles Within Drosophila Larval Segmental Axons

This protocol discusses the live dissection of Drosophila larvae for the purpose of imaging the movement of GFP tagged axonal vesicles on microtubule tracks.

Elucidating the mechanisms of axonal transport has shown to be very important in determining how defects in long distance transport affect different ...

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

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

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

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

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

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date this tool has been widely used to study the regulation of cellular proliferation, differentiation and neuronal migration especially in the developing cerebral cortex 6-8. Here we detail our protocol to electroporate in utero the cerebral cortex and the hippocampus and provide evidence that this approach can be used to study dendrites and spines in these two cerebral regions.
Visualization and manipulation of neurons in primary cultures have contributed to a better understanding of the processes involved in dendrite, spine and synapse development. However neurons growing in vitro are not exposed to all the physiological cues that can affect dendrite and/or spine formation and maintenance during normal development. Our knowledge of dendrite and spine structures in vivo in wild-type or mutant mice comes mostly from observations using the Golgi-Cox method 9. However, Golgi staining is considered to be unpredictable. Indeed, groups of nerve cells and fiber tracts are labeled randomly, with particular areas often appearing completely stained while adjacent areas are devoid of staining. Recent studies have shown that IUE of fluorescent constructs represents an attractive alternative method to study dendrites, spines as well as synapses in mutant / wild-type mice 10-11 (Figure 1A). Moreover in comparison to the generation of mouse knockouts, IUE represents a rapid approach to perform gain and loss of function studies in specific population of cells during a specific time window. In addition, IUE has been successfully used with inducible gene expression or inducible RNAi approaches to refine the temporal control over the expression of a gene or shRNA 12. These advantages of IUE have thus opened new dimensions to study the effect of gene expression/suppression on dendrites and spines not only in specific cerebral structures (Figure 1B) but also at a specific time point of development (Figure 1C).
Finally, IUE provides a useful tool to identify functional interactions between genes involved in dendrite, spine and/or synapse development. Indeed, in contrast to other gene transfer methods such as virus, it is straightforward to combine multiple RNAi or transgenes in the same population of cells. 
In summary, IUE is a powerful method that has already contributed to the characterization of molecular mechanisms underlying brain function and disease and it should also be useful in the study of dendrites and spines.

Visualization and Genetic Manipulation of Dendrites and Spines in the Mouse Cerebral Cortex and Hippocampus using In utero Electroporation

This article describes in detail a protocol to electroporate in utero the cerebral cortex and the hippocampus at E14.5 in mice. We also show that this is a valuable method to study dendrites and spines in these two cerebral regions.

In utero electroporation (IUE) has become a powerful technique to study the development of different regions of the embryonic nervous system 1-5. To date ...

Voltage-sensitive Dye Recording from Axons, Dendrites and Dendritic Spines of Individual Neurons in Brain Slices

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding how the brain works. The primary function of any nerve cell is to process electrical signals, usually from multiple sources. Electrical properties of neuronal processes are extraordinarily complex, dynamic, and, in the general case, impossible to predict in the absence of detailed measurements. To obtain such a measurement one would, ideally, like to be able to monitor, at multiple sites, subthreshold events as they travel from the sites of origin on neuronal processes and summate at particular locations to influence action potential initiation. This goal has not been achieved in any neuron due to technical limitations of measurements that employ electrodes. To overcome this drawback, it is highly desirable to complement the patch-electrode approach with imaging techniques that permit extensive parallel recordings from all parts of a neuron. Here, we describe such a technique - optical recording of membrane potential transients with organic voltage-sensitive dyes (Vm-imaging) - characterized by sub-millisecond and sub-micrometer resolution. Our method is based on pioneering work on voltage-sensitive molecular probes 2. Many aspects of the initial technology have been continuously improved over several decades 3, 5, 11. Additionally, previous work documented two essential characteristics of Vm-imaging. Firstly, fluorescence signals are linearly proportional to membrane potential over the entire physiological range (-100 mV to +100 mV; 10, 14, 16). Secondly, loading neurons with the voltage-sensitive dye used here (JPW 3028) does not have detectable pharmacological effects. The recorded broadening of the spike during dye loading is completely reversible 4, 7. Additionally, experimental evidence shows that it is possible to obtain a significant number (up to hundreds) of recordings prior to any detectable phototoxic effects 4, 6, 12, 13. At present, we take advantage of the superb brightness and stability of a laser light source at near-optimal wavelength to maximize the sensitivity of the Vm-imaging technique. The current sensitivity permits multiple site optical recordings of Vm transients from all parts of a neuron, including axons and axon collaterals, terminal dendritic branches, and individual dendritic spines. The acquired information on signal interactions can be analyzed quantitatively as well as directly visualized in the form of a movie.

An imaging technique for monitoring of membrane potential changes with sub-micrometer spatial and sub-millisecond temporal resolution is described. The technique, based on laser excitation of voltage-sensitive dyes, allows measurements of signals in axons and axon collaterals, terminal dendritic branches, and individual dendritic spines.

Understanding the biophysical properties and functional organization of single neurons and how they process information is fundamental for understanding ...

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 ...

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this information to the brain. As such, determining how olfactory information is transferred to the brain, modifying both perception and behavior, merits investigation. Interestingly, there is emerging evidence that cellular transduction and transcriptional factors are involved in the diversification of olfactory receptor neuron. Here we provide a robust whole mount immunological labeling method to assay in vivo olfactory receptor neuron organization. Using this method, we identified all olfactory receptor neurons with anti-ELAV antibody, a known pan-neural marker and Or49a-mCD8::GFP, an olfactory receptor neuron specifically expressed in Nba neuron using anti-GFP antibody.

Whole Mount Immunolabeling of Olfactory Receptor Neurons in the Drosophila Antenna

Herein we describe the process of whole mount immunostaining of Drosophila antennae, which enables us to better understand the molecular mechanisms involved in the diversification of olfactory receptor neurons (ORN)s.

Odorant molecules bind to their target receptors in a precise and coordinated manner. Each receptor recognizes a specific signal and relays this ...

Production and Isolation of Axons from Sensory Neurons for Biochemical Analysis Using Porous Filters

Neuronal axons use specific mechanisms to mediate extension, maintain integrity, and induce degeneration. An appropriate balance of these events is required to shape functional neuronal circuits. The protocol described here explains how to use cell culture inserts bearing a porous membrane (filter) to obtain large amounts of pure axonal preparations suitable for examination by conventional biochemical or immunocytochemical techniques. The functionality of these filter inserts will be demonstrated with models of developmental pruning and Wallerian degeneration, using explants of embryonic dorsal root ganglion. Axonal integrity and function is compromised in a wide variety of neurodegenerative pathologies. Indeed, it is now clear that axonal dysfunction appears much earlier in the course of the disease than neuronal soma loss in several neurodegenerative diseases, indicating that axonal-specific processes are primarily targeted in these disorders. By obtaining pure axonal samples for analysis by molecular and biochemical techniques, this technique has the potential to shed new light into mechanisms regulating the physiology and pathophysiology of axons. This in turn will have an impact in our understanding of the processes that drive degenerative diseases of the nervous system.

This article describes a neuronal culture system that can be used to obtain large quantities of pure axonal samples for biochemical and immunocytochemical analyses. This technique will allow an improved understanding of normal axonal physiology and signaling pathways leading to neurodegeneration.

Neuronal axons use specific mechanisms to mediate extension, maintain integrity, and induce degeneration. An appropriate balance of these events is ...

Removal of Drosophila Muscle Tissue from Larval Fillets for Immunofluorescence Analysis of Sensory Neurons and Epidermal Cells

Drosophila larval dendritic arborization (da) neurons are a popular model for investigating mechanisms of neuronal morphogenesis. Da neurons develop in communication with the epidermal cells they innervate and thus their analysis benefits from in situ visualization of both neuronally and epidermally expressed proteins by immunofluorescence. Traditional methods of preparing larval fillets for immunofluorescence experiments leave intact the muscle tissue that covers most of the body wall, presenting several challenges to imaging neuronal and epidermal proteins. Here we describe a method for removing muscle tissue from Drosophila larval fillets. This protocol enables imaging of proteins that are otherwise obscured by muscle tissue, improves signal to noise ratio, and facilitates the use of super-resolution microscopy to study da neuron development.

Removal of Drosophila Muscle Tissue from Larval Fillets for Immunofluorescence Analysis of Sensory Neurons and Epidermal Cells

Studies of neuronal morphogenesis using Drosophila larval dendritic arborization (da) neurons benefit from in situ visualization of neuronal and epidermal proteins by immunofluorescence. We describe a procedure that improves immunofluorescence analysis of da neurons and surrounding epidermal cells by removing muscle tissue from the larval body wall.

Drosophila larval dendritic arborization (da) neurons are a popular model for investigating mechanisms of neuronal morphogenesis. Da neurons develop in ...

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 ...

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 ...