SG is supported by funds from the State University of New York at Buffalo and from John R. Oishei Foundation.

1. Preparation of Reagents
<ol>
 <li>Prepare 1x Dissection Buffer using 128 mM NaCl, 4mM CaCl2, 4mM MgCl2, 2 mM KCl, 5mM HEPES and 36mM Sucrose. PH the solution to 7.2 and filter sterilize. </li>
	<li>Prepare the fixative using 1:1 dilution of 16% formaldehyde and 1x Dissection Buffer.</li>
	<li>Prepare 1x PBT using 1x Phosphate Buffered Saline (PBS) at a PH of 7.2 and Triton X-100 in a 1:100 ratio.</li>
	<li>Prepare 5% Bovine Serum Albumin (BSA) with 0.01% Na Azide in 1x PBS.</li>
</ol>
2. Preparation for the Dissection
<ol>
 <li>Sylgard dishes are used for dissection. Prepare the dishes by pouring Sylgard 184 silicone elastomer base and curing agent mixture into a Petri dish. Allow the mixture to completely solidify and dry before using.</li>
	<li>Before the dissection, gather a clean pair of #5 forceps, a clean pair of straight blade vannas scissors, and pins.</li>
</ol>
3. Dissection of 3rd Instar Larvae
<ol>
 <li>Collect wandering 3rd instar larvae of a specific genotype onto a petri dish.</li>
	<li>Wash the larvae in deionized water to get rid of the food.</li>
	<li>Place a larva on the sylgard dish, with the dorsal side up. Pin the larva at its anterior and posterior ends.</li>
	<li>Place a drop of 1x Dissection Buffer on the pinned larva. Using fine scissors nip the larva on the dorsal midline near the posterior end. Using the scissors, cut the larval cuticle from the posterior end through to the anterior end. </li>
	<li>Place a drop of new 1x Dissection Buffer onto the dissected larva. Using a pin, pick one lateral edge of the cuticle near the middle of the larva and pin the cuticle. Using another pin, pin the second lateral edge of the cuticle as shown in the diagram. Two pins are used on each lateral side. See diagram below.</li>
	<li>Carefully remove the intestines, fat bodies, and trachea, leaving only the brain with the two optical lobes and ventral ganglia with the segmental nerves attached. The remainder of the procedure is done on the sylgard dish.</li>
</ol>
4. Fixation of the Dissected Larva
<ol>
	<li>Incubate the dissected larva in fix for 30 minutes, by replacing the fix every 15 minutes.</li>
	<li>After fixation, rinse the larva in 1x PBT for 30 minutes by replacing the buffer every 10 minutes. It is important to note that the larva should always be in a buffer.</li>
</ol>
5. Antibody Staining of the Dissected Larva
<ol>
	<li>Prepare the primary antibody by diluting it to the appropriate concentration in 5% BSA with Na Azide in 1x PBT. Optimal concentrations will differ for different antibodies.</li>
	<li>Replace the last 1x PBT rinse with the prepared primary antibody solution, and incubate in a humid chamber at 4&deg;C overnight. The humid chamber is constructed by lining a plastic dish with kim-wipes soaked in water. </li>
	<li>The following day, rinse the dissected larva in 1x PBT for 30 minutes by replacing the buffer every 10 minutes.</li>
	<li>Prepare the secondary antibody by diluting it to the appropriate concentration in 5% BSA with Na Azide in 1x PBT, and incubate the larva with the secondary antibody solution at 25&deg;C for an hour. Wrap the humid chamber in foil to prevent light, as secondary antibodies are light sensitive.</li>
	<li>After incubation, rinse the larva in 1x PBT for 30 minutes by replacing the buffer every 10 minutes.</li>
</ol>
6. Mounting and Visualization of the Dissected Larva
<ol>
	<li>Before mounting the larva on the slide, prepare the slide by spreading two vertical lines of nail polish in the middle, with a space enough for a cover slip to sit on (see diagram). Let the nail polish dry completely.</li>
	<li>Place a drop of mounting buffer in the space between the vertical nail polish lines on the slide (see diagram). The slide is now ready for mounting. <img src="/files/ftp_upload/2128/2128.6.2.1.jpg" alt="Figure 1 step 6.2" /> <img src="/files/ftp_upload/2128/2128.6.2.2.jpg" alt="Figure 2 step 6.2" /></li>
	<li>After the last 1x PBT rinse, gently remove the lateral pins. Be careful to not tear the cuticle.</li>
	<li>Carefully remove the two remaining pins and pick up the larva with forceps and place the larva horizontally on the prepared slide. Make sure the larva is not flipped over and is oriented ventral side up.</li>
	<li>Gently place a cover slip on top and seal the edges with nail polish (see diagram). The larva is now ready for visualization under a fluorescent microscope. <img src="/files/ftp_upload/2128/2128.6.5.jpg" alt="Figure 1 step 6.5" /></li>
</ol>
7. Representative Results
<img src="/files/ftp_upload/2128/2128fig1.jpg" alt="Figure 1" /> Figure 1: A representative image of larval segmental nerves from a wild type larva using antibodies against the synaptic vesicle protein cysteine string protein (CSP, Zinsmaier et al.1994 ). Note that the segmental nerves are smoothly stained. Bar=20&mu;m
<img src="/files/ftp_upload/2128/2128fig2.jpg" alt="Figure 2" /> Figure 2: A representative image of larval segmental nerves from a motor protein mutant larva using antibodies against the synaptic vesicle protein cysteine string protein (CSP). Note that the segmental nerves show massive accumulations that stain brightly for CSP (arrows).

<ol>
	<li>Gunawardena, S., Goldstein, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disruption+of+Axonal+Transport+and+Neuronal+Viability+by+Amyoid+Precursor+Protein+Mutations+in+Drosophila.">Disruption of Axonal Transport and Neuronal Viability by Amyoid Precursor Protein Mutations in Drosophila.</a> Neuron. 32 (2), 389-401 (2001).</li><li>Gunawardena, S., Her, L. S., Brusch, R. G., Laymon, R. A., Niesman, I. R., Gordesky-Gold, B., Sintasath, L., Bonini, N. M., Goldstein, L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Disruption+of+axonal+transport+by+loss+of+huntingtin+or+expression+of+pathogenic+polyQ+proteins+in+Drosophila.">Disruption of axonal transport by loss of huntingtin or expression of pathogenic polyQ proteins in Drosophila.</a> Neuron. 40 (1), 1-2 (2003).</li><li>Hurd, D. D., Saxton, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinesin+mutations+cause+motor+neuron+disease+phenotypes+by+disrupting+fast+axonal+transport+in+Drosophila.">Kinesin mutations cause motor neuron disease phenotypes by disrupting fast axonal transport in Drosophila.</a> Genetics. 144 (3), 1075-1085 (1996).</li><li>Zinsmaier, K. E., Eberle, K. K., Buchner, E., Walter, N., Benzer, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Paralysis+and+early+death+in+cysteine+string+protein+mutants+of+Drosophila.">Paralysis and early death in cysteine string protein mutants of Drosophila.</a> Science. 263, 977-980 (1994).</li></ol>

No conflicts of interest declared.

The Drosophila larval segmental nerves are a powerful system to study mechanisms in axonal transport. If the 3rd instar larval dissection is performed correctly, all components of the CNS, PNS, and associated tissue, such as the muscles and NMJs, can be examined within a single larva. We have adapted this protocol from the one published by Hurd and Saxton (1996). This protocol can also be adapted to test other neuronal antibodies, but optimization of antibodies should be done. The primary and secondary antibody incubation times can be changed or a blocking step can be added. We have successfully used this protocol to investigate the role of the amyloid precursor protein 1 and Huntingtin 2 in axonal transport.

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		<th>Catalogue Number</th>
		<th>Comment</th>
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<tr>
<td>Dumont #5 Forceps</td>
<td>&nbsp;</td>
<td>Fine Science Tools, Inc.</td>
<td>11252-20</td>
<td>&nbsp;</td>
<tr>
<td>Minutien Pins, Stainless Steel</td>
<td>&nbsp;</td>
<td>Fine Science Tools, Inc.</td>
<td>26002-10</td>
<td>&nbsp;</td>
<tr>
<td>Mcpherson-Vannas Straight Blade 8cm Scissors</td>
<td>&nbsp;</td>
<td>Fisher Scientific</td>
<td>50822236</td>
<td>&nbsp;</td>
<tr>
<td>Vectashield, Mounting Medium</td>
<td>&nbsp;</td>
<td>Vector Laboratories, Inc.</td>
<td>H-1000</td>
<td>&nbsp;</td>
<tr>
<td>Sylgard 184 Silicone elastomer</td>
<td>&nbsp;</td>
<td>Dow Corning Corporation</td>
<td>4026148</td>
<td>&nbsp;</td>
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<td>formaldehyde, 16%</td>
<td>&nbsp;</td>
<td>Electron Microscopy Sciences</td>
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<td>&nbsp;</td>
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<td>&nbsp;</td>
<td>Developmental Studies Hybridoma Bank</td>
<td>&nbsp;</td>
<td>&nbsp;</td>
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<td>Alexa Fluor 568 Goat Anti-Mouse IgG</td>
<td>&nbsp;</td>
<td>Invitrogen</td>
<td>A-11004</td>
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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.

Watch this Scientific Journal Video about Visualization of Larval Segmental Nerves in 3rd Instar Drosophila Larval Preparations at JoVE.com

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.

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 ...

visualization-larval-segmental-nerves-3rd-instar-drosophila-larval

Nanyang Technological University

Research

JoVE Journal

Neuroscience

10.3K Views.  SUNY-University at Buffalo. 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.

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

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 ...

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 ...

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).

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.

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

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).

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.

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

Forebrain Electrophysiological Recording in Larval Zebrafish

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked seizures, epilepsy can be acquired as a result of an insult to the brain or a genetic mutation. Efforts to model seizures in animals have primarily utilized acquired insults (convulsant drugs, stimulation or brain injury) and genetic manipulations (antisense knockdown, homologous recombination or transgenesis) in rodents. Zebrafish are a vertebrate model system1-3 that could provide a valuable alternative to rodent-based epilepsy research. Zebrafish are used extensively in the study of vertebrate genetics or development, exhibit a high degree of genetic similarity to mammals and express homologs for ~85% of known human single-gene epilepsy mutations. Because of their small size (4-6 mm in length), zebrafish larvae can be maintained in fluid volumes as low as 100 &mu;l during early development and arrayed in multi-well plates. Reagents can be added directly to the solution in which embryos develop, simplifying drug administration and enabling rapid in vivo screening of test compounds4. Synthetic oligonucleotides (morpholinos), mutagenesis, zinc finger nuclease and transgenic approaches can be used to rapidly generate gene knockdown or mutation in zebrafish5-7. These properties afford zebrafish studies an unprecedented statistical power analysis advantage over rodents in the study of neurological disorders such as epilepsy. Because the "gold standard" for epilepsy research is to monitor and analyze the abnormal electrical discharges that originate in a central brain structure (i.e., seizures), a method to efficiently record brain activity in larval zebrafish is described here. This method is an adaptation of conventional extracellular recording techniques and allows for stable long-term monitoring of brain activity in intact zebrafish larvae. Sample recordings are shown for acute seizures induced by bath application of convulsant drugs and spontaneous seizures recorded in a genetically modified fish.

A simple method to record extracellular field potentials in the larval zebrafish forebrain is described. The method provides a robust in vivo read-out of seizure-like activity. This technique can be used with genetically modified zebrafish larvae carrying epilepsy-related genes or seizures evoked by administration of convulsant drugs.

Epilepsy affects nearly 3 million people in the United States and up to 50 million people worldwide. Defined as the occurrence of spontaneous unprovoked ...

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 ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

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 ...

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic pathways. Quantitative descriptions are rarely performed, although genetic manipulations produce a range of phenotypic effects and variations are observed even among individuals within control groups.&#160;Emerging evidence shows that morphology, size and location of organelles play a previously underappreciated, yet fundamental role in cell function and survival. Here we provide step-by-step instructions for performing quantitative analyses of phenotypes at the Drosophila larval neuromuscular junction (NMJ). We use several reliable immuno-histochemical markers combined with bio-imaging techniques and morphometric analyses to examine the effects of genetic mutations on specific cellular processes. In particular, we focus on the quantitative analysis of phenotypes affecting morphology, size and position of nuclei within the striated muscles of Drosophila larvae. The Drosophila larval NMJ is a valuable experimental model to investigate the molecular mechanisms underlying the structure and the function of the neuromuscular system, both in health and disease. However, the methodologies we describe here can be extended to other systems as well.

Why Quantification Matters: Characterization of Phenotypes at the Drosophila Larval Neuromuscular Junction

Morphology, size and location of intracellular organelles are evolutionarily conserved and appear to directly affect their function. Understanding the molecular mechanisms underlying these processes has become an important goal of modern biology. Here we show how these studies can be facilitated by the application of quantitative techniques.

Most studies on morphogenesis rely on qualitative descriptions of how anatomical traits are affected by the disruption of specific genes and genetic ...

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 ...